Methods for making proteins containing free cysteine residues

ABSTRACT

The present invention relates to novel methods of making soluble proteins having free cysteines in which a host cell is exposed to a cysteine blocking agent. The soluble proteins produced by the methods can then be modified to increase their effectiveness. Such modifications include attaching a PEG moiety to form pegylated proteins.

FIELD OF THE INVENTION

The present invention relates generally to methods of making proteinsand more specifically to recombinant proteins containing a “free”cysteine residue that does not form a disulfide bond.

BACKGROUND OF THE INVENTION

There is considerable interest on the part of patients and healthcareprovides in the development of low cost, long-acting, “user-friendly”protein therapeutics. Proteins are expensive to manufacture and unlikeconventional small molecule drugs, are usually not readily absorbed bythe body. Moreover they are digested if taken orally. Therefore,proteins must typically be administered by injection. After injectionmost proteins are cleared rapidly from the body, necessitating frequent,often daily, injections. Patients dislike injections, which leads toreduced compliance and reduced drug efficacy. Some proteins such aserythropoietin (EPO) are effective when administered less often (threetimes per week for EPO) but this is due to the fact that the proteinsare glycosylated. Glycosylation requires that the recombinant proteinsbe man using mammalian cell expression systems, which is expensive andincreases the cost of protein pharmaceuticals.

Thus, there is a strong need to develop protein delivery technologiesthat lower the costs of protein therapeutics to patients and healthcareproviders. One solution to this problem is the development of methods toprolong the circulating half-lives of protein therapeutics in the bodyso that the proteins do not have to be injected frequently. Thissolution also satisfies the needs and desires of patients for proteintherapeutics that are “user-friendly”, i.e., protein therapeutics thatdo not require frequent injections.

Covalent modification of proteins with polyethylene glycol (PEG) hasproven to be a useful method to extend the circulating half-lives ofproteins in the body (Abuchowski et al. 1984; Hershfield, 1987; Meyerset al., 1991). Covalent attachment of PEG to a protein increases theprotein's effective size and reduces its rate of clearance from thebody. PEGs are commercially available in several sizes, allowing thecirculating half-lives of PEG-modified proteins to be tailored forindividual indications through use of different size PEGs. Otherdocumented in vivo benefits of PEG modification are an increase inprotein solubility, stability (possibly due to profusion of the proteinfrom proteases) and a decrease in protein immunogenicity (Katre et al.,1987; Katre, 1990).

One known method for PEGylating proteins uses compounds such asN-hydroxy succinimide (NHS)-PEG to attach PEG to free amines, typicallyat lysine residues or at the N-terminal amino acid. A major limitationof this approach is that proteins typically contain several lysines, inaddition to the N-terminal amino acid, and the PEG moiety attaches tothe protein non-specifically at any of the available free amines,resulting in a heterogeneous product mixture. Many NHS-PEGylatedproteins are unsuitable for commercial use because of low specificactivities and heterogeneity. Inactivation results from covalentmodification of one or more lysine residues or the N-terminal amino acidrequired for biological activity or from covalent attachment of the PEGmoiety near the active site of the protein.

Of particular relevance to this application is the finding thatmodification of human growth hormone (hGH) with amine-reactive reagents,including NHS-PEG reagents, reduces biological activity of the proteinby more than 10-fold (Teh and Chapman, 1988; Clark et al., 1996). GH isa 22 kDa protein secreted by the pituitary gland. GH stimulatesmetabolism of bone, cartilage and muscle and is the body's primaryhormone for stimulating somatic growth during childhood. Recombinanthuman GH (rhGH) is used to treat short stature resulting from GHinadequacy, Turner's Syndrome and renal failure in children. GH is notglycosylated and is fully active when produced in bacteria. The proteinhas a short in vivo half-life and must be administered by dailysubcutaneous injection for maximum effectiveness (McGillivray et al.,1996).

There is considerable interest in the development of long-acting formsof hGH. Attempts to create long-acting forms of hGH by PEGylating theprotein with amine-reactive PEG reagents have met with limited successdue to significant reductions in bioactivity upon PEGylation. Further,the protein becomes PEGylated at multiple sites (Clark et al., 1996).hGH contains nine lysines, in addition to the N-terminal amino acid.Certain of these lysines are located in regions of the protein known tobe critical for receptor binding (Cunningham et al., 1989; Cunninghamand Wells, 1989). Modification of these lysine residues significantlyreduces receptor binding and bioactivity of the protein (de la Llosa etal., 1985; Martal et al., 1985; Teh and Chapman, 1988; Cunningham andWells, 1989). hGH is readily modified by NHS-PEG reagents, butbiological activity of the NHS-PEG protein is severely compromised,amounting to only 1% of wild type GH biological activity for a GHprotein modified with five 5 kDa PEG molecules (Clark et al., 1996). TheEC₅₀ for this multiply PEGylated GH protein is 440 ng/ml orapproximately 20 nM (Clark et al., 1996). In addition to possessingsignificantly reduced biological activity, NHS-PEG-hGH is veryheterogeneous due to different numbers of PEG molecules attached to theprotein and at different amino acid residues, which has an impact on itsusefulness as a potential therapeutic. Clark et al. (1996) showed thatthe circulating half-life of NHS-PEG-hGH in animals is significantlyprolonged relative to non-modified GH. Despite possessing significantlyreduced in vitro biological activity, NHS-PEG-hGH was effective andcould be administered less often than non-modified hGH in a ratGH-deficiency model (Clark et al., 1996). However, high doses ofNHS-PEG-hGH (60-180 μg per injection per rat) were required for efficacyin the animal models due to the low specific activity of the modifiedprotein. There is a clear need for better methods to create PEGylatedhGH proteins that retain greater bioactivity. There also is a need todevelop methods for PEGylating hGH in a way that creates a homogeneousPEG-hGH product.

Biological activities of several other commercially important proteinsare significantly reduced by amine-reactive PEG reagents. EPO containsseveral lysine residues that are critical for bioactivity of the protein(Boissel et al., 1993; Matthews et al., 1996) and modification of lysineresidues in EPO results in near complete loss of biological activity(Wojchowski and Caslake, 1989). Covalent modification ofalpha-interferon-2 with amine-reactive PEGs results in 40-75% loss ofbioactivity (Goodson and Katre, 1990; Karasiewicz et al., 1995). Loss ofbiological activity is greatest with large (e.g., 10 kDa) PEGs(Karasiewicz et al., 1995). Covalent modification of GCSF withamine-reactive PEGs results in greater than 60% loss of bioactivity(Tanaka et al., 1991). Extensive modification of IL-2 withamine-reactive PEGs results in greater than 90% loss of bioactivity(Goodson and Katre, 1990).

A second known method for PEGylating proteins covalently attaches PEG tocysteine residues using cysteine-reactive PEGs. A number of highlyspecific, cysteine-reactive PEGs with different reactive groups (e.g.,maleimide, vinylsulfone) and different size PEGs (240 kDa) arecommercially available. At neutral pH, these PEG reagents selectivelyattach to “free,” cysteine residues, i.e., cysteine residues notinvolved in disulfide bonds. Cysteine residues in most proteinsparticipate in disulfide bonds and are not available for PEGylationusing cysteine-reactive PEGs. Through in vitro mutagenesis usingrecombinant DNA techniques, additional cysteine residues can beintroduced anywhere into the protein. The newly added “free” cysteinescan serve as sites for the specific attachment of a PEG molecule usingcysteine-reactive PEGs. The added cysteine residue can be a substitutionfor an existing amino acid in a protein, added preceding theamino-terminus of the protein or after the carboxy-terminus of theprotein, or inserted between two amino acids in the protein.Alternatively, one of two cysteines involved in a native disulfide bondmay be deleted or substituted with another amino acid, leaving a nativecysteine (the cysteine residue in the protein that normally would form adisulfide bond with the deleted or substituted cysteine residue) freeand available for chemical modification. Preferably the amino acidsubstituted for the cysteine would be a neutral amino acid such asserine or alanine. Growth hormone has two disulfide bonds that can bereduced and alkylated with iodoacetimide without impairing biologicalactivity (Bewley et al., (1969). Each of the four cysteines would bereasonable targets for deletion or substitution by another amino acid.

Several naturally-occurring proteins are known to contain one or more“free” cysteine residues. Examples of such naturally-occurring proteinsinclude human Interleukin (IL)-2, beta interferon (Mark et al., 1984),G-CSF (Lu et al., 1989) and basic fibroblast growth factor (Thompson,1992). IL-2, G-CSF and beta interferon contain an odd number of cysteineresidues, where as basic fibroblast growth factor contains an evennumber of cysteine residues.

However, expression of recombinant proteins containing free cysteineresidues has been problematic due to reactivity of the free sulfhydrylat physiological conditions. Several recombinant proteins containingfree cysteines have been expressed as intracellular proteins in bacteriasuch as E. coli. Examples include natural proteins such as IL-2, betainterferon, G-CSF, basic fibroblast growth factor and engineeredcysteine muteins of IL-2 (Goodson and Katre, 1990), IL-3 (Shaw et al.,1992), Tumor Necrosis Factor Binding Protein (Tuma et al., 1995), IGF-I(Cox and McDermott, 1994), IGFBP-1 (Van Den Berg et al., 1997) andprotease nexin and related proteins (Braxton, 1998). All of theseproteins were insoluble when expressed intracellularly in bacteria. Theinsoluble proteins could be refolded into their native conformations byperforming a series of denaturation, reduction and refolding procedures.These steps add time and cost to the manufacturing process for producingthe proteins in bacteria. Improved stability and yields of IL-2 (Mark etal., 1985) and beta interferon (DeChiara et al., 1986) have beenobtained by substituting another amino acid, e.g., serine, for the freecysteine residue. It would be preferable to express the recombinantproteins in a soluble, biologically active form to eliminate these extrasteps.

One known method of expressing soluble recombinant proteins in bacteriais to secrete them into the periplasmic space or into the media. It isknown that certain recombinant proteins such as GH are expressed in asoluble active form when they are secreted into the E. coli periplasm,whereas they are insoluble when expressed intracellularly in E. coli.Secretion is achieved by fusing DNA sequences encoding growth hormone orother proteins of interest to DNA sequences encoding bacterial signalsequences such as those derived from the stII (Fujimoto et al., 1988)and ompA proteins (Ghrayeb et al., 1984). Secretion of recombinantproteins in bacteria is desirable because the natural N-terminus of therecombinant protein can be maintained. Intracellular expression ofrecombinant proteins requires that an N-terminal methionine be presentat the amino-terminus of the recombinant protein. Methionine is notnormally present at the amino-terminus of the mature forms of many humanproteins. For example, the amino-terminal amino acid of the mature formof human growth hormone is phenylalanine. An amino-terminal methioninemust be added to the amino-terminus of a recombinant protein, if amethionine is not present at this position, in order for the protein tobe expressed efficiently in bacteria. Typically addition of theamino-terminal methionine is accomplished by adding an ATG methioninecodon preceding the DNA sequence encoding the recombinant protein. Theadded N-terminal methionine often is not removed from the recombinantprotein, particularly if the recombinant protein is insoluble. Such isthe case with hGH, where the N-terminal methionine is not removed whenthe protein is expressed intracellularly in E. coli. The addedN-terminal methionine creates a “non-natural” protein that potentiallycan stimulate an immune response in a human. In contrast, there is noadded methionine on hGH that is secreted into the periplasmic spaceusing stII (Chang et al., 1987) or ompA (Cheah et al., 1994) signalsequences; the recombinant protein begins with the native amino-terminalamino acid phenylalanine. The native hGH protein sequence is maintainedbecause of bacterial enzymes that cleave the std-hGH protein (orompA-hGH protein) between the stII (or ompA) signal sequence and thestart of the mature hGH protein. While the periplasmic space is believedto be an oxidizing environment that should promote disulfide bondformation, coexpression of protein disulfide isomerase with bovinepancreatic trypsin inhibitor resulted in a six-fold increase in theyield of correctly folded protein from the E. coli periplasm (Ostermeieret al., (1996). This result would suggest that periplasmic proteinfolding can at times be inefficient and is in need of improvement forlarge scale protein production.

hGH has four cysteines that form two disulfides. hGH can be secretedinto the E. coli periplasm using stII or ompA signal sequences. Thesecreted protein is soluble and biologically active (Hsiung et al.,1986). The predominant secreted form of hGH is a monomer with anapparent molecular weight by sodium dodecyl sulfate polyacrylamide gelelectrophoresis (SDS-PAGE) of 22 kDa Recombinant hGH can be isolatedfrom the periplasmic space by using an osmotic shock procedure (Koshlandand Botstein, 1980), which preferentially releases periplasmic, but notintracellular, proteins into the osmotic shock buffer. The released hGHprotein is then purified by column chromatography ((Hsiung et al.,1986).

When similar procedures were attempted to secrete hGH variantscontaining a free cysteine residue (five cysteines; 2N+1), it wasdiscovered that the recombinant hGH variants formed multimers andaggregates when isolated using standard osmotic shock and purificationprocedures developed for hGH. Very little of the monomeric hGH variantproteins could be detected by non-reduced SDS-PAGE in the osmotic shocklysates or during purification of the proteins by column chromatography.

Alpha interferon (IFN-α2) also contains four cysteine residues that formtwo disulfide bonds. IFN-α2 can be secreted into the E. coli periplasmusing the stII signal sequence (Voss et al., 1994). The secreted proteinis soluble and biologically active (Voss et al., 1994). The predominantsecreted form of IFN-α2 is a monomer with an apparent molecular weightby SDS-PAGE of 19 kDa Secreted recombinant IFN-α2 can be purified bycolumn chromatography (Voss et al., 1994).

When similar procedures were attempted to secrete IFN-α2 variantscontaining a free cysteine residue (five cysteines; 2N+1), it wasdiscovered that the recombinant IFN-α2 variants formed multimers andaggregates when isolated using standard purification proceduresdeveloped for IFN-α2. The IFN-α2 variants eluted from the columns verydifferently than IFN-α2 and very little of the monomeric IFN-α2 variantproteins could be purified using column chromatography proceduresdeveloped for IFN-α2.

An alternative method to synthesizing a protein containing a freecysteine residue is to introduce a thiol group into a proteinpost-translationally via a chemical reaction with succinimidyl6-[3-2-pyridyldithio)propionamido]hexanoate (LC-SPDP, commerciallyavailable from Pierce Chemical Company). LC-SPDP reacts with lysineresidues to create a free sulfhydryl group. Chemically cross-linkeddimeric EPO was prepared using this reagent in conjunction with amaleimide protein modifying reagent (Sytkowsk et al., 1998). Aheterologous mixture of chemically cross-linked EPO proteins wasrecovered after purification due to non-specific modification of thevarious lysine residues in EPO. Enhanced pharmacokinetics and in vivopotency of the chemically cross-linked EPO proteins were observed.

Another method that has been used to increase the size of a protein andimprove its in vivo potency involves dimerization of the protein usingchemical crosslinking reagents. GH is thought to transduce a cellularsignal by cross-linking two GH receptors. A GH-GH dimer might facilitateenhanced receptor dimerization and subsequent amplification of theintracellular signal.

Chemically cross-linked dimeric hGH proteins have been described byMockridge et al. (1998). Using a water soluble cross-linking reagent1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide (EDC), GH was randomlyderivatized to give predominantly amide-linked dimers but alsoamide-linked multimers, depending on the concentration of EDC reagentused. While an increase in in vivo potency was observed, the finalprotein preparation was heterogeneous due to non-specific reaction ofthe EDC reagent with various amino acids in the protein, includinglysine, aspartic acid and glutamic acid residues and the amino- andcarboxy-termini. Injection of such a preparation into humans would beundesirable due to the toxic nature of EDC, potential immunogenicresponse to the unnatural amide bond formed between the proteins.Generating consistent batches of a purified protein also would bedifficult at the manufacturing scale.

Therefore, despite considerable effort, a need still exists for aprocess for generating homogeneous preparations of long actingrecombinant proteins by enhancement of protein molecular weight. A needalso for methods that allow secretion and recovery of recombinantproteins containing free cysteine residues in high yield. The presentinvention satisfies these needs and provides related advantages as well.

SUMMARY OF THE INVENTION

The present invention relates to methods for obtaining a soluble proteinhaving a free cysteine. The methods are generally accomplished byobtaining a host cell capable of expressing the soluble protein,exposing the host cell to a cysteine blocking agent, and isolating thesoluble protein from the host cell. In one embodiment in which theprotein is not secreted into the media by th host cell, the host cell isdisrupted in the presence of the cysteine blocking agent and the solubleprotein is isolated or purified from the soluble fraction of thedisrupted host cell. In another embodiment in which the soluble proteinis secreted by the host cell into the media, the host cell is exposed tothe cysteine blocking agent before, during or after synthesis of thesoluble protein by the host cell

Suitable host cells include bacteria, yeast, insect or mammalian cells.Preferably, the host cell is a bacterial cell, particularly E. coli.

Preferably, the soluble protein produced by the methods of the presentinvention are recombinant proteins, especially cysteine variants ormuteins of a protein. The methods are useful for producing proteinsincluding, without limitation, human growth hormone, EPO and interferon,especially alpha interferon, their derivatives or antagonists. Otherproteins include members of the TGF-beta superfamily, platelet derivedgrowth factor-A, platelet derived growth factor-B, nerve growth factor,brain derived neurotophic factor, neurotrophin-3, neurotrophin-4,vascular endothelial growth factor, or a derivative or an antagonistthereof. Cysteine muteins of heavy or light chain of an immunoglobulinor a derivative thereof are also contemplated.

Useful cysteine blocking agents include any thiol-reactive compound,including for example, cystine, cystamine, dithioglycolic acid, oxidizedglutathione, iodine, hydrogen peroxide, dihydroascorbic acid,tetrathionate, O-iodosobenzoate or oxygen in the presence of a metalion.

The present methods further include various methods of attaching a PEGmoiety to the soluble protein to form pegylated proteins in which thePEG moiety is attached to the soluble protein through the free cysteine.Higher order multimeric proteins involving the coupling of two or moreof the soluble proteins are also within the present invention.

The present invention further includes the soluble proteins and theirderivatives, including pegylated proteins, made by the methods disclosedherein. Such pegylated proteins include monopegylated hGh, EPO and alphainterferon.

The present invention also provides methods for pegylating the solubleproteins obtained by the methods described herein. Such methods includepurifying the protein, reducing at least partially the protein with adisulfide-reducing agent and exposing the protein to a cysteine-reactivemoiety. Optionally, the modified cysteine protein can be isolated fromunmodified protein.

Methods of treating conditions treatable by growth hormone, EPO andalpha interferon are also within the present invention. The solubleproteins or their derivatives, including pegylated derivatives, areadministered to patients suffering from conditions in which known growthhormone, EPO or alpha interferon is effective.

BRIEF DESCRIPTION OF THE FIGURE

FIG. 1 is a diagram of mutagenesis by overlap extension in which twoseparate fragments are amplified from a target DNA segment.

DESCRIPTION OF THE INVENTION

The present invention provides novel methods of obtaining proteinshaving free cysteine residues. The invention further provides novelproteins, particularly recombinant proteins, produced by these novelmethods as well as derivatives of such recombinant proteins. The novelmethods for preparing such proteins are generally accomplished by:

-   -   (a) obtaining a host cell capable of expressing a protein having        a free cysteine;    -   (b) exposing the host cell to a cysteine blocking agent; and    -   (c) isolating the protein from the host cell.

In one embodiment, the methods include the steps of disrupting the hostcell in the presence of the cysteine blocking agent followed byisolating the protein from the soluble fraction of the disrupted cell.

In a further embodiment in which the proteins are secreted into themedia by prokaryotic or eukaryotic host cells, the methods include thesteps of:

-   -   (a) obtaining a host cell capable of expressing a protein having        a free cysteine;    -   (b) exposing the host cell to a cysteine blocking agent during        synthesis, or after synthesis but prior to purification, of the        protein having a free cysteine residue; and    -   (d) isolating the protein from other cellular components in the        media.

As identified above, the first step in these methods is to obtain a hostcell capable of expressing a protein having a free cysteine residue.Suitable host cells can be prokaryotic or eukaryotic. Examples ofappropriate host cells that can be used to express recombinant proteinsinclude bacteria, yeast, insect and mammalian cells. Bacteria cells areparticularly useful especially E. coli.

As used herein, the term “protein having a flee cysteine residue” meansany natural or recombinant protein-peptide containing 2N+1 cysteineresidues, where N can be 0 or any integer, and proteins or peptidescontaining 2N cysteines, where two or more of the cysteines do notnormally participate in a disulfide bond. Thus, the methods of thepresent invention are useful in enhancing the expression, recovery andpurification of any protein or peptide having a free cysteine,particularly cysteine added variant recombinant proteins (referred toherein as “cysteine muteins” or “cysteine variants”) having one or morefree cysteines and/or having 2 or more cysteines that naturally form adisulfide bond. Although the expression, recovery and purification of anatural protein having a free cysteine expressed by its natural hostcell can be enhanced by the methods of the present invention, thedescription herein predominantly refers to recombinant proteins forillustrative purposes only. In addition, the proteins can be derivedfrom any anima species including human, companion animals and farmanimals.

Accordingly, the present invention encompasses a wide variety ofrecombinant proteins. These proteins include, but are not limited to,glial-derived neurotrophic factor (GDNF), transforming growthfactor-beta1 (TGF-beta1), TGF-beta2, TGF-beta3, inhibin A, inhibin B,bone morphogenetic protein-2 (BMP-2), BMP-4, inhibin alpha, Mullerianinhibiting substance (MIS), OP-1 (osteogenic protein 1), which are allmembers of the TGF-beta superfamily. The monomer subunits of theTGF-beta superfamily share certain structural features: they generallycontain 8 highly conserved cysteine residues that form 4 intramoleculardisulfides. Typically a ninth conserved cysteine is free in themonomeric form of the protein but participates in an intermoleculardisulfide bond formed during the homodimerization or heterodimericationof the monomer subunits. Other members of the TGF-beta superfamily aredescribed by Massague (1990), Daopin et al. (1992), Kingsley (1994),Kutty et al. (1998), and Lawton et al. (1997), incorporated herein byreference.

Immunoglobulin heavy and light chain monomers also contain cysteineresidues that participate in intramolecular disulfides as well as freecysteines (Roitt et al., 1989 and Paul, 1989). These free cysteinesnormally only participate in disulfide bonds as a consequence ofmultimerization events such as heavy chain homodimerization, heavychain-light chain heterodimerization, homodimerization of the (heavychain-light chain) heterodimers, and other higher order assemblies suchas pentamerization of the (heavy chain-light chain) heterodimers in thecase of IgM. Thus, the methods of the present invention can be employedto enhance the expression, recovery and purification of heavy and/orlight chains (or various domains thereof) of human immunoglobulins suchas IgG1, IgG2, IgG3, IgG4, IgM IgA1, IgA2, secretory IgA, IgD and IgE.Immunoglobulins from other species could also be similarly expressed,recovered and purified. Proteins genetically fused to immunoglobulins orimmunoglobulin domains as described in Chamow & Ashkenazi (1996) couldalso be similarly expressed, recovered and purified.

The present methods can also enhance the expression, recovery andpurification of additional recombinant proteins including members ofgrowth hormone superfamily. The following proteins are encoded by genesof the growth hormone (GH) supergene family (Bazan (1990); Bazan (1991);Mott and Campbell (1995); Silvennoinen and Ihle (1996); Martinet al.,1990; Hannum et al., 1994): growth hormone, prolactin, placentallactogen, erythropoietin (EPO), thrombopoietin (TPO), interleukin-2(IL-2), IL-3, L-4, IL-5, IL-6, IL-7, IL-9, IL-10, IL-il, EL-12 (p35subunit), IL-13, IL-15, oncostatin M, ciliary neurotrophic factor,leukemia inhibitory factor, alpha interferon, beta interferon, gammainterferon, omega interferon, tau interferon, granulocyte-colonystimulating factor (G-CSF), granulocyte-macrophage colony stimulatingfactor (GM-CSF), macrophage colony stimulating factor (M-CSF),cardiotrophin-1 (CT-1), Stem Cell Factor and the flt3/flk2 ligand (“theGH supergene family”). It is anticipated that additional members of thisgene family will be identified in the future through gene cloning andsequencing. Members of the GH supergene family have similar secondaryand tertiary structures, despite the fact that they generally havelimited amino acid or DNA sequence identity. The shared structuralfeatures allow new members of the gene family to be readily identified.

A group of proteins has been classed as a structural superfamily basedon the shared structural motif termed the “cystine knot”. The cystineknot is defined by six conserved cysteine residues that form threeintramolecular disulfide bonds that are topologically “knotted”(McDonald and Hendrickson, 1993). These proteins also form homo- orheterodimers and in some but not all instances dimerization involvesintermolecular disulfide formation. Members of this family include themembers of the TGF-beta superfamily and other proteins such as plateletderived growth factor-A (PDGF-A), PDGF-B, nerve growth factor (NGF),brain derived neurotrophic factor (BDNF), neurotrophin-3 (NT-3), NT-4,and vascular endothelial growth factor (VEGF). Cystine and othercysteine blocking reagents could also enhance expression, recovery andpurification of proteins with this structural motif.

The present methods can also enhance the expression, recovery andpurification of other recombinant proteins and/or cysteine addedvariants of those proteins. Classes of proteins would include proteasesand other enzymes, protease inhibitors, cytokines, cytokine antagonists,allergens, chemokines, gonadotrophins, chemotactins, lipid-bindingproteins, pituitary hormones, growth factors, somatomedans,immunoglobulins, interleukins, interferons, soluble receptors, vaccines,and hemoglobins. Specific examples of proteins include, for example,leptin, insulin, insulin-like growth factor 1 (IGF1), superoxidedismutase, catalase, asparaginse, uricase, fibroblast growth factors,arginase, phenylalanine ammonia, angiostatin, endostatin, Factor VIII,Factor IX interleukin 1 receptor antagonist, protease nexin andanti-thrombin III.

Other protein variants that would benefit from PEGylation and wouldtherefore be reasonable candidates for cysteine added modificationsinclude proteins or peptides with poor solubility or a tendency toaggregate, proteins or peptides that are susceptable to proteolysis,proteins or peptides needing improved mechanical stability, proteins orpeptides that are cleared rapidly from the body, or proteins or peptideswith undesirable immunogenic or antigentic properties.

If desired, muteins of natural proteins can be generally constructedusing site directed PCR-based mutagenesis as decribed in general inMethods in Molecular Biology, Vol. 15: PCR Protocols: Current Methodsand Applications edited by White, B. A. (1993) Humana Press, Inc.,Totowa, N.J. and PCR Protocols: A Guide to Methods and Applicationsedited by Innis, M. A. et al. (1990) Academic Press, Inc. San Diego,Calif. Typically, PCR primer oligonucleotides are designed toincorporate nucleotide changes to the coding sequence of proteins thatresult in substitution of a cysteine residue for an amino acid at aspecific position within the protein. Such mutagenic oligonucleotideprimers can also be designed to incorporate an additional cysteineresidue at the carboxy terminus or amino terminus of the coding sequenceof proteins. In this latter case one or more additional amino acidresidues could also be incorporated amino terminal and/or carboxyterminal to the added cysteine residue if that were desirable. Moreoveroligonucleotides can be designed to incorporate cysteine residues asinsertion mutations at specific positions within the protein codingsequence if that were desirable. Again, one or more additional aminoacids could be inserted along with the cysteine residue and these aminoacids could be positioned amino terminal and/or carboxy terminal to thecysteine residue.

The choice of sequences for mutagenic oligos is dictated by the positionwhere the desired cysteine residue is to be placed and the propinquityof useful restriction endonuclease sites. Generally it is desirable toplace the mutation, i.e. the mismatched segment near the middle of theoligo to enhance the annealing of the oligo to the template. It is alsodesirable for the mutagenic oligo to span a unique restriction site sothat the PCR product can be cleaved to generate a fragment that can bereadily cloned into a suitable vector. An example would be one that canbe used to express the mutein or that provides convenient restrictionsites for excising the mutated gene and readily cloning it into such anexpression vector. It is generally desirable to employ mutagenic oligosunder 80 bases in length and lengths of 30-40 bases are more preferable.Sometimes mutation sites and restriction sites are separated bydistances that are greater than that which is desirable for synthesis ofsynthetic oligonucleotides. In such instances, multiple rounds of PCRcan be employed to incrementally extend the length of the PCR productsuch that it includes the desired useful restriction site or genestargeted for mutagenesis can be reengineered or resynthesized toincorporate restriction sites at appropriate positions. Alternatively,variations of PCR mutagenesis protocols, such as the so-called“Megaprimer Method” (Barik, S. pp277-286 in Methods in MolecularBiology, Vol. 15: PCR Protocols: Current Methods and Applications editedby White, B. A. (1993) Humana Press, Inc., Totowa, N.J.) or “GeneSplicing by Overlap Extension” (Horton, R. M. pp251-261 in Methods inMolecular Biology, Vol. 15: PCR Protocols: Current Methods andApplications edited by White, B. A, (1993) Humana Press, Inc., Totowa,N.J.), both incorporated herein by reference, can also be employed toconstruct such mutations.

Next, the host cell is exposed to a cysteine blocking agent. In oneembodiment, the blocking agent is present at the time of celldisruption, and preferably is added prior to disrupting the cells. Celldisruption can be accomplished by, for example, mechanical sheer such asa French pressure cell, enzymatic digestion, sonication, homogenization,glass bead vortexing, detergent treatment, organic solvents, freezethaw, grinding with alumina or sand and the like (Bollag et al., 1996).

In an alternative embodiment, the cysteine blocking agent can be exposedto the host cell before, during or after the host cell is induced toexpress the desired protein. For example, the host cell can be culturedin the presence of the cysteine blocking agent, which might be preferredfor expression of proteins that are secreted into the media such aserythropoietin, for example. Alternatively, prior to or at the time of,exposing the host cell to the cysteine blocking agent, the host cell canbe induced to express the desired protein, either as a secreted proteinin the periplasm or media, or as a cytoplasmic protein. Methods known inthe art can be used to induce such expression in the cytoplasm or todirect secretion depending on cell origin, including, for example, themethods described in the examples below. A wide variety of signalpeptides have been used successfully to transport proteins to theperiplasmic space. Examples of these include prokaryotic signalsequences such as ompA, stII, PhoA signal (Denefle et al., 1989), OmpT(Johnson et al., 1996), LamB and OmpF (Hoffman and Wright, 1985),beta-lactamase (Kadonaga et al., 1984), enterotoxins LT-A, LT-B(Morioka-Fujimoto et al., 1991), and protein A from S. aureus (Abrahmsenet al., 1986). A number of non-natural, synthetic, signal sequences thatfacilitate secretion of certain proteins are also known to those skilledin the art.

For proteins secreted to the periplasm, an osmotic shock treatment canbe used to selectively disrupt the outer membrane of the host cells withthe resulting release of periplasmic proteins. The osmotic shock buffercan be any known in the art, including, for example, the osmotic shockbuffer and procedures described in Hsiung et al. (1986), or as describedin the examples below. For proteins secreted into the media, preferablythe media should contain the cysteine blocking agent during the time thecells express and secrete the protein. The cysteine blocking agent alsocould be added to the media following secretion of the protein but priorto purification of the protein.

The cysteine blocking agent can be added to the culture media at aconcentration in the range of about 0.1 μM to about 100 mM. Preferably,the concentration of cysteine blocking agent in the media is about 50 μMto about 5 mM

Although not wishing to be bound by any particular theory, it isbelieved that the cysteine blocking agents used in the present methodscovalently attach to the “free” cysteine residue, forming a mixeddisulfide, thus stabilizing the free cysteine residue and preventingmultimerization and aggregation of the protein. Alternatively, thepresence of an oxidizing agent in the osmotic shock buffers may beaugmenting the protein refold process in case incomplete renaturationhad occurred following secretion of the protein into the periplasmicspace. However, as noted above, periplasmic protein refolding can beinefficient. For this reason it is believed that addition of cystine toosmotic shock buffers also may increase the recovery of recombinantproteins containing an even number of cysteine residues, even if thecysteines normally form disulfide bonds. In addition, the inventorsbelieve it may be advantageous to add cystine or similar compounds tothe fermentation media during bacterial growth because these compoundsshould diffuse into the periplasm due to the porous nature of thebacterial outer membrane. Protein folding and blocking of the freecysteine can be accomplished before cell recovery and lysis. Earlyprotein stabilization protects against proteolysis and can contribute tohigher recovery yields of recombinant proteins.

A number of thiol-reactive compounds can be used as cysteine blockingagents to stabilize proteins containing free cysteines. In addition tocystine, blocking agents can also include reagents containing disulfidelinkages such as cystamine, dithioglycolic acid, oxidized glutathione,5,5′-dithiobis(2-nitrobenzoic acid (Ellman's reagent), pyridinedisulfides, compounds of the type R—S—S—CO—OCH₃, other derivatives ofcystine such as diformylcystine, diacetylcystine, diglycylcystine,dialanylcystine diglutaminylcystine, cystinyldiglycine,cystnyldiglutamine, dialanylcystine dianhydride, cystinephenylbydantoin, homocystine, dithiodipropionic acid, dimethylcystine,or any dithiol or chemical capable of undergoing a disulfide exchangereaction. Sulfenyl halides can also be used to prepare mixed disulfides.Other thiol blocking agents that may find use in stabilizing cysteineadded protein variants include compounds that are able to reversiblyreact with free thiols. These agents include certain heavy metals saltsor organic derivatives of zinc, mercury, and silver. Other mercaptideforming agents or reversible thiol reactive compounds are described byR. Cecil and J. R. McPhee (1959) and Torchinskii (1971).

In the final step of the general method, the desired protein isrecovered and purified from the soluble cytoplasmic fraction, thesoluble periplasmic fraction or the soluble fraction of the media Anymethod for recovering and purifying proteins from the media, cytoplasmicor periplasmic fraction can be used. Such recovery and purificationmethods are known or readily determined by those skilled in the art,including, for example, centrifugation, filtration, dialysis,chromatography, including size exclusion, procedures and the like. Asuitable method for the recovery and purification of a desired proteinwill depend, in part, on the properties of the protein and the intendeduse.

The present invention further provides novel methods for producingsoluble interferon, particularly alpha interferon, that results in asignificant increase in the percent of the recovered interferon that hasbeen properly processed. These methods include culturing host cellscapable of expressing interferon at a pH range of about 5 to about 6.5,and preferably about 5.5 to about 6.5. Published reports (Voss et al.(1994)) using a higher pH only resulted in 50% properly processedinterferon, whereas the new methods of the present invention at lowerpHs recovered about 80-90%.

A discovery was made that certain of the monomeric hGH cysteine variantsformed disulfide-linked dimeric hGH proteins during the chromatographyprocedures used to purify these proteins. The disulfide-linked hGHdimers formed when cystine was removed from the column buffers. Newprocedures were developed to purify the disulfide-linked dimeric hGHproteins because the disulfide-linked dimeric hGH proteins behaveddifferently than monomeric hGH proteins during the column chromatographysteps used to purify the proteins. Unexpectedly, it was discovered thatthe disulfide-linked dimeric hGH proteins were biologically active in invitro bioassays. Biologically active, homogeneous, disulfide-linkeddimeric hGH proteins are novel. Accordingly, the present inventionfurther relates to these biologically active, homogeneous di-sulfidelined, dimeric hGH proteins as well. Higher order multimers are alsocontemplated, including trimers, tetramers and the like, as described inthe examples below.

The purified proteins can then be further processed if desired. Forexample, the proteins can be PEGylated at the free cysteine site withvarious cysteine-reactive PEG reagents, and subsequently purified asmonoPEGylated proteins. The term “monoPEGylated” is defined to mean aprotein modified by covalent attachment of a single PEG molecule at aspecific site on the protein. Any method known to those skilled in theart can be used, including, for example, the methods described in theexamples below, particularly Example 11.

Braxton (1998) teaches methods for PEGylating cysteine muteins ofproteins, and in particular cysteine muteins of GH and erythropoietin.Braxton (1998) specifically teaches that the buffers used to PEGylatethe cysteine muteins should not contain a reducing agent. Examples ofreducing agents provided by Braxton (1998) are beta-mercaptoethanol(BME) and dithiothreitol DTT). When similar procedures were used toPEGylate cysteine muteins of GH, erythropoietin and alpha interferon, itwas discovered that the cysteine muteins did not PEGylate. It has nowbeen discovered that treatment of the purified cysteine muteins with areducing agent is required for the proteins to be PEGylated. Althoughnot wanting to be bound by any particular theory, the inventors believethat the reducing agent is required to reduce the mixed disulfide andexpose the free cysteine residue in the protein so that the freecysteine can react with the PEG reagent. Thus, the present inventionalso relates to methods for PEGylating cysteine muteins of GH,erythropoietin, alpha interferon and other proteins containing 2N+1cysteine residues, proteins containing 2N cysteine residues where two ormore of the cysteine residues are free, particularly those muteins andproteins in which the free cysteine residue is blocked by a mixeddisulfide.

The present invention further relates to purified, monoPEGylated proteinvariants produced by the methods disclosed herein that are not onlybiologically active, but also retain high specific activity inprotein-dependent mammalian cell proliferation assays. Such proteinvariants include, for example, the following purified, monPEGylatedcysteine muteins: hGH, EPO and alpha IFN. For example, the in vitrobiological activities of the monoPEGylated hGH variants were 10- to100-fold greater than the biological activity of hGH that has beenPEGylated using NHS-PEG reagents.

In one embodiment of the monoPEGylated hGH, the polyethylene glycol isattached to the C-D loop of hGH and the resulting monoPEGylated hGH hasan EC less than about 110 ng/ml (5 nM), preferably less than about 50ng/ml (2.3 nM). Alternatively, the polyethylene glycol moiety can beattached to a region proximal to the Helix A of hGH and the resultingmonoPEGylated hGH has an EC₅₀ less than about 110 ng/ml (5 nM),preferably less than 11 ng/ml (0.5 nM), and more preferably less thanabout 2.2 ng/ml (0.1 nM).

In one embodiment of the monoPEGylated EPO, the polyethylene glycol isattached to the C-D loop of EPO and the resulting monoPEGylated EPO hasan EC₅₀ less than about 1000 ng/ml (21 nM), preferably less than about100 ng/ml (approximately 6 nM), more preferably less than about 10 ng/ml(approximately 0.6 nM) and most preferably less than about 1 ng/ml(approximately 0.06 nM). Alternatively, the polyethylene glycol moietycan be attached to the A-B loop of EPO and the resulting monoPEGylatedEPO has an EC₅₀ less than about 100 ng/ml (approximately 5 nM),preferably less than 20 ng/ml (approximately 1 nM), and more preferablyless than about 1 ng/ml (approximately 0.05 nM).

In one embodiment of the monoPEGylated alpha IFN, the polyethyleneglycol is attached to the region proximal to Helix A of alpha IFN andthe resulting monoPEGylated alpha IFN has an IC₅₀ less than about 100ng/ml (approximately 5 pM), more preferably less than about 50 pg/ml(approximately 2.5 pM) and most preferably about 22 ng/ml (approximately1.2 pM). Alternatively, the polyethylene glycol moiety can be attachedto the C-D loop of IFN-α2 and the resulting monoPEGylated IFN-α2 has anEC₅₀ less than about 100 pg/ml (approximately 5 pM).

There are over 25 distinct IFN-α genes (Pestka et al., 1987). Members ofthe IFN-α family share varying degrees of amino acid homology andexhibit overlapping sets of biological activities. Non-naturalrecombinant IFN-αs, created through joining together regions ofdifferent IFN-α proteins are in various stages of clinical development(Horisberger and DiMarco, 1995). A non-natural “consensus” interferon(Blatt et al., 1996), which incorporates the most common amino acid ateach position of IFN-α, also has been described. Appropriate sites forPEGylating cysteine muteins of IFN-α2 should be directly applicable toother members of the IFN-α gene family and to non-natural IFN-αs.Kinstler et al., (1996) described monoPEGylated consensus interferon inwhich the protein is preferentially mono PEGylated at the N-terminal,non-natural methionine residue. Bioactivity of the PEGylated protein wasreduced approximately 5-fold relative to non-modified consensusinterferon (Kinstler et al., 1996).

The present invention further provides protein variants that can becovalently attached or conjugated to each other or to a chemical groupto produce higher order multimers, such as dimers, trimers andtetramers. Such higher order multimers can be produced according tomethods known to those skilled in the art or as described in Example 15below. For example, such a conjugation can produce a hGH, EPO or alphaIFN adduct having a greater molecular weight than the correspondingnative protein. Chemical groups suitable for coupling are preferablynon-toxic and non-immunogenic. These chemical groups would includecarbohydrates or polymers such as polyols.

The “PEG moiety” useful for attaching to the cysteine variants of thepresent invention to form “pegylated” proteins include any suitablepolymer, for example, a linear or branched chained polyol. A preferredpolyol is polyethylene glycol, which is a synthetic polymer composed ofethylene oxide units. The ethylene oxide units can vary such thatPEGylated-protein variants can be obtained with apparent molecularweights by size-exclusion chromatography ranging from approximately30-500,000. The size of the PEG moiety dirty impacts its circulatinghalf-life Yamaoka et al. (1994). Accordingly, one could engineer proteinvariants with differing circulating half-lives for specific therapeuticapplications or preferred dosing regimes by varying the size orstructure of the PEG moiety. Thus, the present invention encompasses GHprotein variants having an apparent molecular weight greater than about30 kDa, and more preferably greater than about 70 kDa as determined bysize exclusion chromatography, with an EC₅₀ less than about 400 ng/ml(18 nM), preferably less than 10 ng/ml (5 nM), more preferably less thanabout 10 ng/ml (0.5 nM), and even more preferably less than about 2.2ng/ml (0.1 nM). The present invention further encompasses EPO proteinvariants having an apparent molecular weight greater than about 30 kDa,and more preferably greater than about 70 kDa as determined by sizeexclusion chromatography, with an EC₅₀ less than about 1000 ng/ml (21nM), preferably less than 100 ng/ml (6 nM), more preferably less thanabout 10 ng/ml (0.6 nM), and even more preferably less than about 1ng/ml (0.06 nM). The present invention further encompasses alpha IFN(IFN-α) protein variants having an apparent molecular weight greaterthan about 30 kDa, and more preferably greater than about 70 kDa asdetermined by size exclusion chromatography, with an IC₅₀ less thanabout 1900 pg/ml (100 pM), preferably less than 400 pg/ml (21 pM), morepreferably less than 100 pg/ml (5 nM), and even more preferably lessthan about 38 pg/ml (2 pM).

The reactive PEG end group for cysteine modification includes but is notlimited to vinylsulfone, maleimide and iodoacetyl moieties. The PEG endgroup should be specific for free thiols with the reaction occurringunder conditions that are not detrimental to the protein.

Antagonist hGH variants also can be prepared using a cysteine-addedvariant GH where chemical derivatization does not interfere withreceptor binding but does prohibit the signaling process. Conditionsthat would benefit from the administration of a GH antagonist includeacromegaly, vascular eye diseases, diabetic nephropathy, restenosisfollowing angioplasty and growth hormone responsive malignancies.

As used herein, the term “derivative” refers to any variant of a proteinexpressed and recovered by the present methods. Such variants include,but are not limited to, PEGylated versions, dimers and other higherorder variants, amino acid variants, fusion proteins, changes incarbohydrate, phosphorylation or other attached groups found on naturalproteins, and any other variants disclosed herein.

The compounds produced by the present methods can be used for a varietyof in vitro and in vivo uses. The proteins and their derivatives of thepresent invention can be used for research, diagnostic or therapeuticpurposes that are known for their wildtype, natural, or previously-knownmodified counterparts. In vitro uses include, for example, the use ofthe protein for screening, detecting and/or purifying other proteins.

For therapeutic uses, one skilled in the art can readily determine theappropriate dose, frequency of dosing and route of administration.Factors in making such determinations include, without limitation, thenature of the protein to be administered, the condition to be treated,potential patient compliance, the age and weight of the patient, and thelike. The compounds of the present invention can also be used asdelivery vehicles for enhancement of the circulating half-life of thetherapeutics that are attached or for directing delivery to a specifictarget within the body.

The following examples are not intended to be limiting, but onlyexemplary of specific embodiments of the invention.

EXAMPLE 1 Development of an In Vitro Bioassay for Human Growth Hormone

An hGH cell proliferation assay that uses the murine FDC-P1 cell linestably transfected with the rabbit GH receptor was developed (Rowlinsonet al., 1995). The mouse FDC-P1 cell line was purchased from theAmerican Type Culture Collection and routinely propagated in RPMI 1640media supplemented with 10% fetal calf serum, 50 μg/ml penicillin, 50μg/ml streptomycin, 2 mM glutamine and 17-170 Units/ml mouse IL-3(FDC-P1 media).

A. Cloning a cDNA Encoding the Rabbit GH Receptor

The rabbit GH receptor was cloned by PCR using forward primer BB3(5′-CCCCGGATCCGCCACCATGGATCTCTGG CAGCTGCTGTT-3′) (SEQ.ID.NO. 1) andreverse primer BB36 (5′-CCCCGTCGACTCTAGAGCCATTAGATACAAAGCTCT TGGG-3′)(SEQ.ID.NO. 2). BB3 anneals to the DNA sequence encoding the initiatormethionine and amino terminal portion of the receptor. BB3 contains anoptimized KOZAK sequence preceding the initiator methionine and a Bam HIsite for cloning purposes. BB36 anneals to the 3′ untranslated region ofthe rabbit GH receptor mRNA and contains Xba I and Sal I restrictionsites for cloning purposes. Rabbit liver poly(A)⁺ mRNA (purchased fromCLONTECH, Inc.) was used as the substrate in first strand synthesis ofsingle-stranded cDNA to produce template for PCR amplification. Firststrand synthesis of single-stranded cDNA was accomplished using a 1stStrand cDNA Synthesis Kit for RT-PCR (AMV) from Boehringer MannheimCorp. Parallel first strand cDNA syntheses were performed using randomhexamers or BB36 as the primer. Subsequent PCR reactions using theproducts of the first strand syntheses as templates were carried outwith primers BB3 and BB36. The expected ˜1.9 kb PCR product was observedin PCR reactions using random hexamer-primed or BB36-primed cDNA astemplate. The random hexamer-primed cDNA was digested with Bam HI andXba I, which generates two fragments (˜365 bp and ˜1600 bp) because therabbit GH receptor gene contains an internal Bam HI site. Both fragmentswere gel-purified. The full-length rabbit GH receptor cDNA was thencloned in two steps. First the ˜1600 bp Bam HI-Xba I fragment was clonedinto pcDNA3.1(+) (Invitrogen Corporation) that had been digested withthese same two enzymes. These clones were readily obtained at reasonablefrequencies and showed no evidence of deletions as determined byrestriction digests and subsequent sequencing. To complete the rabbitreceptor cDNA clone, one of the sequenced plasmids containing the 1600bp Bam HI-Xba I fragment was digested with Bam HI, treated with calfalkaline phosphatase, gel-purified and ligated with the gel-purified˜365 bp Bam HI fragment that contains the 5′ portion of the rabbit GHreceptor gene. Transformants from this ligation were picked and analyzedby restriction digestion and PCR to confirm the presence of the ˜365 bpfragment and to determine its orientation relative to the distal segmentof the rabbit GH receptor gene. The sequence for one full length clonewas then verified. This plasmid, designated pBBT118, was used to stablytransfect FDC-P1 cells.

B. Selection of Stably Transfected FDC-P1 Cells Expressing the Rabbit GHReceptor

Endotoxin-free pBBT118 DNA was prepared using a Qiagen “Endo-FreePlasmid Purification Kit” and used to transfect FDC-P1 cells. Mouse IL-3was purchased from R&D Systems. FDC-P1 cells were transfected withplasmid pBBT118 using DMRIE-C cationized lipid reagent purchased fromGIBCO, following the manufacturer's recommended directions. Briefly, 4μg of plasmid DNA were complexed with 4-30 μl of the DMRIE-C reagent in1 ml of OptiMEM media (GIBCO) for 45 minutes in six well tissue culturedishes. Following complex formation, 2×10⁶ FDC-P1 cells in 200 μl ofOptiMEM media supplemented with mouse IL-3 were added to each well andthe mixture left for 4 h at 37° C. The final mouse IL-3 concentrationwas 17 Units/ml. Two ml of FDC-P1 media containing 15% fetal bovineserum were added to each well and the cells left overnight at 37° C. Thenext day transfected cells were transferred to T-75 tissue cultureflasks containing 15 ml FDC-P1 media supplemented with IL-3 (17 U/ml),hGH (5 nM) and 10% horse serum rather than fetal calf serum. Horse serumwas used because of reports that fetal calf serum contains agrowth-promoting activity for FDC-P1 cells. Three days later the cellswere centrifuged and resuspended in fresh FDC-P1 media containing 400μg/ml G418, 17 U/ml L-3, 5 nM hGH, 10% horse serum and incubated at 37°C. Media was changed every few days. The cells from each transfectionwere split into T-75 tissue culture flasks containing fresh media andeither mouse IL-3 (17 U/ml) or hGH (5 nM). G418 resistant cells wereobtained from both the IL-3- and hGH-containing flasks. Thetransformants used in the bioassays originated from flasks containinghGH. Twelve independent cell lines were selected by limiting dilution.Five of the cell lines (GH-R3, -R4, -R5, -R6 and -R9) showed a goodproliferative response to hGH. Preliminary experiments indicated thatthe EC₅₀ for hGH was similar for each cell line, although the magnitudeof the growth response varied depending upon the line. The GH-R4 cellline was studied in most detail and was used for the assays presentedbelow. The cell lines were routinely propagated in RPMI 1640 mediacontaining 10% horse serum, 50 Units/ml penicillin, 50 μg/mlstreptomycin, 2 mM glutamine, 400 μg/ml G418 and 2-5 nM pituitary hGH orrhGH.

C. Development of an hGH Bioassay Using FDC-P1 Expressing the Rabbit GHReceptor

A modified version of the MTT cell proliferation assay described byRowlinson et al. (1995) was developed to measure hGH bioactivity. Ourassay measures uptake and reduction of the dye MTS, which creates asoluble product, rather than MTT, which creates an insoluble productthat must be solubilized with organic solvents. The advantage of usingMTS is that absorbance of the wells can be determined without the needto lyse the cells with organic solvents.

GH-R4 cells were washed three times with phenol red-free RPMI 1640 mediaand suspended in assay media (phenol red-free RPMI 1640, 10% horseserum, 50 Units/ml penicillin, 50 μg/ml streptomycin, 400 μg/ml G418) ata concentration of 1×10⁵ cells/ml. Fifty microliters of the cellsuspension (5×10³ cells) were added to wells of a 96 well flat bottomedtissue culture plate. Serial three-fold dilutions of protein sampleswere prepared in assay media and added to microtiter wells in a volumeof 50 μl, yielding a final volume of 100 μl per well. Protein sampleswere assayed in triplicate wells. Plates were incubated at 37° C. in ahumidified 5% CO₂ tissue culture incubator for 66-72 h, at which time 20μl of an MTS/PES (PES is an electron coupler) reagent mixture (CellTiter96 Aqueous One solution reagent, Promega Corporation) was added to eachwell. Absorbance of the wells at 490 nm was measured 2-4 h later.Absorbance value means +/−standard deviations for the triplicate wellswere calculated. Control wells contained media but no cells. Absorbancevalues for the control wells (typically 0.06-0.2 absorbance units) weresubtracted from the absorbance values for the sample wells. Controlexperiments demonstrated that absorbance signals correlated with cellnumber up to absorbance values of 2. All assays included a humanpituitary GH standard. Experiments utilizing the parental FDC-P1 cellline were performed as described above except that the assay media didnot contain G418 and fetal calf serum was substituted for horse serum.

EXAMPLE 2 Cloning and Expression of rhGH

A. Cloning a cDNA encoding human Growth Hormone (GH)

A human GH cDNA was amplified from human pituitary single-stranded cDNA(commercially available from CLONTECH, Inc., Palo Alto, Calif.) usingthe polymerase chain reaction (PCR) technique and primers BB1 and BB2.The sequence of BB1 is(5′-GGGGGTCGACCATATGTTCCCAACCATTCCCTTATCCAG-3′)(SEQ.ID.NO. 3). Thesequence of BB2 is (5′-GGGGGATCCTCACTAGAAGCCACAGCTGCCCTC-3′)(SEQ.ID.NO.4). Primer BB 1 was designed to encode an initiator methionine precedingthe first amino acid of mature GH, phenylalalanine, and Sal I and Nde Isites for cloning purposes. The reverse primer, BB2, contains a Bam HIsite for cloning purposes. The PCR reactions contained 20 pmoles of eacholigo primer, 1×PCR buffer (Perkin-Elmer buffer containing MgCl₂), 200μM concentration of each of the four nucleotides dA, dC, dG and dT, 2 ngof single-stranded cDNA, 2.5 units of Taq polymerase (Perkin-Elmer) and2.5 units of Pfu polymerase (Stratagene, Inc). The PCR reactionconditions were: 96° C. for 3 minutes, 35 cycles of (95° C., 1 minute;63° C. for 30 seconds; 72° C. for 1 minute), followed by 10 minutes at72° C. The thermocycler employed was the Amplitron II Thermal Cycler(Thermolyne) The approximate 600 bp PCR product was digested with Sal Iand Bam HI gel-purified and cloned into similarly digested plasmid pUC19(commercially available from New England BioLabs, Beverly, Mass.). Theligation mixture was transformed into E. coli strain DH5alpha andtransformants selected on LB plates containing ampicillin. Severalcolonies were grown overnight in LB media and plasmid DNA isolated usingminiplasmid DNA isolation kits purchased from Qiagen, Inc (Valencia,Calif.). Clone LB6 was determined to have the correct DNA sequence.

For expression in E. coli, clone LB6 was digested with Nde I and Eco RI,the approximate 600 bp fragment gel-purified, and cloned into plasmidpCYB1 (commercially available from New England Biolabs, Beverly, Mass.)that had been digested with the same enzymes and treated with calfalkaline phosphatase. The ligation mixture was transformed into E. coliDH5alpha and transformants selected on LB ampicillin plates. Plasmid DNAwas isolated from several tmnsformants and screened by digestion withNde I and Eco RI. A correct clone was identified and named pCYB1: wtGH(pBBT120). This plasmid was transformed into E. coli strains JM109 orW3110 (available from New England BioLabs and the American Type CultureCollection).

B. Construction of stII-GH

Wild type GH clone LB6 (pUC19: wild type GH) was used as the template toconstruct a GH clone containing the E. coli stII signal sequence.Because of its length, the stII sequence was added in two sequential PCRreactions. The first reaction used forward primer BB12 and reverseprimer BB10. BB10 has the sequence (5′CGCGGATCCGATTAGAATCCACAGCTCCCCTC3′)(SEQ.ID.NO. 5). BB12 has the sequence(5′-GCATCTATGTTCGTTTTCTCTATCGCTACCAACGCTTACGCATTCCCAACCATTCCCTTATCCAG-3′)(SEQ.ID.NO.6). The PCR reactions were as described for amplifying wild type GH. Theapproximate 630 bp PCR product was gel-purified using the Qiaex II GelExtraction Kit (Qiagen, Inc), diluted 50-fold in water and 2 μl used astemplate for the second PCR reaction. The second PCR reaction usedreverse primer BB10 and forward primer BB11. BB11 has the sequence(5′CCCCCTCTAGACATATGAAGAAGAACATCGCATTCCTGCTGGCATCTATGTTCGTTTTCTCTATCG-3′)(SEQ.ID.NO.7). Primer BB11 contains XbaI and NdeI sites for cloning purposes. PCRconditions were as described for the first reaction. The approximate 660bp PCR product was digested with XbaI and BamHI, gel-purified and clonedinto similarly cut plasmid pcDNA3.1(+) (Invitrogen, Inc. Carlsbad,Calif.). Clone pCDNA3.1(+)::stII-GH(5C) or “5C” was determined to havethe correct DNA sequence.

Clone “5C” was cleaved with NdeI and BamHI and cloned into similarly cutpBBT108 (a derivative of pUC19 which lacks a Pst I site, this plasmid isdescribed below). A clone with the correct insert was identifiedfollowing digestion with these enzymes. This clone, designated pBBT111,was digested with Nde I and Sal I, the 660 bp fragment containing thestII-GH fusion gene, was gel-purified and cloned into the plasmidexpression vector pCYB1 (New England BioLabs) that had been digestedwith the same enzymes and treated with calf alkaline phosphatase. Arecombinant plasmid containing the stII-GH insertion was identified byrestriction endonuclease digestions. One such isolate was chosen forfurther studies and was designated pBBT114. This plasmid was transformedinto E. coli stains JM109 or W3110 (available from New England BioLabsand the American Type Culture Collection).

C. Construction of ompA-GH

Wild type GH clone LB6 (pUC19: wild type GH) was used as the template toconstruct a GH clone containing the E. coli ompA signal sequence.Because of its length, the ompA sequence was added in two sequential PCRreactions. The first reaction used forward primer BB7(5′GCAGTGGCACTGGCTGGTTTCGCTACCGTAGCGCAGGCCTTCCCAACCATTCCCTTATCCAG3′)(SEQ.ID.NO. 8) and reverse primer BB10:(5′CGCGGATCCGATRAGAATCCACAGCTCCCCTC 3′)(SEQ.ID.NO. 5). The PCR reactionswere as described for amplifying wild type GH except that approximately4 ng of plasmid LB6 was used as the template rather than single-strandedcDNA and the PCR conditions were 96° C. for 3 minutes, 30 cycles of (95°C. for 1 minute; 63° C. for 30 seconds; 72° C. for 1 minute) followed by72° C. for 10 minutes. The approximate 630 bp PCR product wasgel-purified using the Qiaex II Gel, Extraction Kit (Qiagen, Inc),diluted 50-fold in water and 2 μl used as template for the second PCRreaction. The second PCR reaction used reverse primer BB10 and forwardprimer BB6: (5′CCCCGTCGACACATATGAAGAAGACAGCTATCGCGATTGCAGTGGCACTGGCTGGTTTC 3′)(SEQ.ID.NO. 9). PCR conditions were as described forthe first reaction. The approximate 660 bp PCR product was gel-purified,digested with Sal I and Bam HI and cloned into pUC19 (New EnglandBioLabs) which was cut with Sal I and Bam HI or pcDNA3.1(+) (Invitrogen)which had been cut by Xho I and Bam HI (Sal I and Xho I producecompatible single-stranded overhangs). When several clones weresequenced, it was discovered that all pUC19 (8/8) clones containederrors in the region of the ompA sequence. Only one pcDNA3.1(+) clonewas sequenced and it contained a sequence ambiguity in the ompA region.In order to generate a correct ompA-GH fusion, gene segments of twosequenced clones, which contained different errors separated by aconvenient restriction site were recombined and cloned into thepUC19-derivative that lacks the Pst I site (see pBBT108 describedbelow). The resulting plasmid, termed pBBT112, carries the ompA-GHfusion gene cloned as an Nde I-Bam HI fragment into these same sites inpBBT108. This plasmid is designated pBBT112 and is used in PCR-based,site-specific mutagenesis of GH as described below.

D. Construction of Pst I⁻pUC19 (pBBT 108)

To facilitate mutagenesis of the cloned GH gene for construction ofselected cysteine substitution and insertion mutations, a derivative ofthe plasmid pUC19 (New England BioLabs) lacking a Pst I site wasconstructed as follows. pUC19 plasmid DNA was digested with Pst I andsubsequently treated at 75° C. with Pfu DNA Polymerase (Stratagene)using the vendor-supplied reaction buffer supplemented with 200 μMdNTPs. Under these conditions the polymerase will digest the 3′single-stranded overhang created by Pst I digestion but will not digestinto the double-stranded region and the net result will be the deletionof the 4 single-stranded bases which comprise the middle four bases ofthe Pst I recognition site. The resulting molecule has double-stranded,i.e. “blunt”, ends. Following these enzymatic reactions the linearmonomer was gel-purified using the Qiaex II Gel Extraction Kit (Qiagen,Inc). This purified DNA was treated with T4 DNA Ligase (New EnglandBioLabs) according to the vendor protocols, digested with Pst I, andused to transform E. coli DH5alpha. Transformants were picked andanalyzed by restriction digestion with Pst I and Bam HI. One of thetransformants which was not cleaved by Pst I but was cleaved at thenearby Bam HI site was picked and designated pBBT108.

E. Expression of met-hGH and stII-hGH in E. coli

pBBT120, which encodes met-hGH, and pBBT114, which encodes stII-hGH weretransformed into E. coli strain W3110. The parental vector pCYB1 alsowas transformed into W3110. The resulting strains were given thefollowing designations:

-   -   BOB130: W3110(pCYB1)=vector only    -   BOB133: W3110(pBBT120)=met-hGH    -   BOB132: W3110(pBBT114)=stII-hGH

These strains were grown overnight at 37° C. in Luria Broth (LB)containing 100 μg/ml ampicillin. The saturated overnight cultures werediluted to ˜0.025 O.D.s at A₆₀₀ in LB containing 100 μg/ml ampicillinand incubated at 37° C. in shake flasks. When culture O.D.s reached0.25-0.5, IPTG was added to a final concentration of 0.5 mM to induceexpression of the recombinant proteins. For initial experiments,cultures were sampled at 0, 1, 3, 5 and ˜16 h post-induction. Samples ofinduced and uninduced cultures were pelleted and resuspended in SDS-PAGEsample buffer with the addition of 1% β-mercaptoethanol (BME) whendesirable. Samples were electrophoresed on precast 14% Tris-glycinepolyacrylamide gels. Gels were stained with Coomassie Blue or wereanalyzed by Western blotting.

Coomassie staining of whole cell lysates from strains BOB133, expressingmet-hGH, and BOB132, expressing stII-hGH, showed a band of ˜22 kDa thatco-migrated with a purified rhGH standard purchased from ResearchDiagnostics Inc. The rhGH band was most prominent in induced culturesfollowing overnight induction. Western blots using a polyclonal rabbitanti-hGH antiserum purchased from United States Biological, Inc.confirmed the presence of rhGH in lysates of induced cultures of BOB132and BOB133 at both 3 and 16 h post-induction No rhGH as detected byWestern Blotting of induced cultures of the control strain BOB130(vector only) at 3 or 16 h post-induction.

An induced culture of BOB132 (expressing stII-hGH) was prepared asdescribed above and subjected to osmotic shock based on the analyticalprocedure of Koshland and Botstein (1980). This procedure ruptures theE. coli outer membrane and releases the contents of the periplasm intothe surrounding medium. Subsequent centrifugation separates the solubleperiplasmic components (recovered in the supernatant) from cytoplasmic,insoluble periplasmic, and cell-associated components (recovered in thepellet). Specifically, E. coli strain W3110 containing the s-II hGHplasmid was grown at 37° C. overnight in LB containing 100 μg/mlampicillin. The saturated overnight culture was diluted to 0.03 O.D. atA₆₀₀ in 25 ml of LB containing 100 μg/ml ampicillin and incubated at 37°C. in a 250 ml shake flask. When the culture O.D. reached approximately0.4, 100 mM IPTG was added for a final concentration of 0.5 mM to induceexpression of the recombinant protein. The induced culture was thenincubated at 37° C. overnight (˜16 h). The induced overnight culturereached an O.D. of 3.3 at A₆₀₀ and 4 O.D.s (1.2 ml) was centrifuged in aEppendorf model 5415C microfuge at 14,000 rpm for 5 minutes at 4° C. Thecell pellet was resuspended to approximately 10 O.D. in ice cold 20%sucrose, 10 mM Tris-HCl pH 8.0 by trituration and vortexing and EDTA pH8.0 was added for a final concentration of 17 mM. Resuspended cells wereincubated on ice for 10 minutes and centrifuged as above. The resultantpellet was resuspended at 10 O.D.s in ice cold water by trituration andvortexing and incubated on ice for 10 minutes. The resuspended cellswere then centrifuged as above and the resultant supernatant (solubleperiplasmic fraction) and cell pellet (insoluble periplasmic and cellassociated components) were analyzed by SDS-PAGE.

The bulk of the rhGH synthesized by BOB132 was found to be soluble andlocalized to the periplasm. The periplasmic rhGH protein wasindistinguishable in size from a purified pituitary hGH standardindicating that the stII signal sequence was removed during proteinsecretion.

EXAMPLE 3 Purification and Bioactivity of rhGH

A. Purification of Wild-Type rhGH

In order to purify a significant quantity of wild-type rhGH, a 330 mlculture of BOB132 (expressing stII-hGH) was induced, cultured overnightand subjected to osmotic shock based on the preparative proceduredescribed by Hsiung et al. (1986). Specifically, E. coli strain W3110containing the st-II hGH plasmid was grown at 37° C. overnight in LBcontaining 100 μg/ml ampicillin. The saturated overnight culture wasdiluted to 0.03 O.D. at A₆₀₀ in 2×250 ml (500 ml total volume) of LBcontaining 100 μg/ml ampicillin and incubated at 37° C. in 2 L shakeflasks. When the culture's O.D. reached approximately 0.4, 100 mM IPTGwas added for a final concentration of 0.5 mM to induce expression ofthe recombinant protein. The induced culture was incubated at 37° C.overnight (˜16 h). The induced overnight cultures reached an O.D. ofapproximately 3.8 at A₆₀₀ and were centrifuged using a Sorval RC-5centrifuge and a GSA rotor at 8,000 rpm for 5 minutes at 4° C. The cellpellets were combined and resuspended to approximately 47 O.D. in icecold 20% sucrose, 10 mM Tris-HCl pH 8.0 by trituration and EDTA pH 8.0was added for a final concentration of 25 mM. Resuspended cells wereincubated on ice for 10 minutes and centrifuged in an IEC Centra MP4Rcentrifuge with an 854 rotor at 8,500 rpm for 7 minutes at 4° C. Theresultant pellets were resuspended at 47 O.D. in ice cold water bytrituration and vortexing and incubated on ice for 30 minutes. Theresuspended cells were centrifuged in the IEC centrifuge as above andthe resultant supernatant (soluble periplasmic fraction) and cell pellet(insoluble periplasmic and cell associated components) were analyzed bySDS-PAGE. Again the gel showed that rhGH produced was soluble,periplasmic, and indistinguishable in size from the pitutary hGHstandard.

rhGH was purified in a two step procedure based on that described byBecker and Hsiung (1986). The supernatant from the osmotic shock wasloaded onto a 5 ml Pharmacia HiTrap Q Sepharose column equilibrated in10 mM Tris-HCl pH 8.0 and the bound proteins were eluted with a 15column volume 50-250 mM linear NaCl gradient Column fractions wereanalyzed by SDS-PAGE. Fractions 22-25 eluting at a salt concentration ofaround 100-125 mM were enriched for hGH, and were pooled, concentratedand further fractionated on a Superdex 200 HR 10/30 sizing column.Fractions 34-36 from the Superdex column (representing MWs around 21-22kDA based on the elution profile of MW standards) contained most of therhGH, were pooled and stored as frozen aliquots at 0° C. The final yieldof rhGH, as determined by absorbance at 280 nm and by using a Bradfordprotein assay kit (Bio-Rad Laboratories), was about 2 mg. Non-reducedpituitary hGH migrates with a slightly smaller apparent molecular weightthan reduced pituitary hGH when analyze by SDS-PAGE. This molecularweight change is indicative of proper disulfide bond formation Ourpurified rhGH co-migrated with pituitary hGH under both reducing andnon-reducing conditions indicating that the rhGH was properly folded anddisulfide-bonded. Data presented below indicates that the biologicalactivity of rhGH is indistinguishable from that of pituitary hGH.

B. Bioassay Results for Pituitary hGH and rhGH

The parental FDC-P1 cell line shows a strong proliferative response tomouse IL-3, but not to pituitary hGH. In the absence of IL-3, themajority of FDC-P1 cells die, giving absorbance values less than 0.2. Incontrast, FDC-P1 cells transformed with the rabbit growth hormonereceptor proliferate in response to pituitary hGH, as evidenced by adose-dependent increase in cell number and absorbance values. The EC₅₀(protein concentration required to achieve half-maximal stimulation) forthis effect ranged from 0.75-1.2 ng/ml pituitary hGH (0.03-0.05 nM) indifferent experiments, similar to what has been reported in theliterature (Rowlinson et al., 1995). A significant difference betweenthe parental FDC-PL line and FDC-P1 cells transformed with the rabbitgrowth hormone receptor is that the latter cells survive in the absenceof IL-3 or hGH, resulting in higher absorbance values (typically0.6-1.1, depending upon the assay and length of incubation with MTS inthe zero growth factor control wells). The initial pool of rabbit growthhormone receptor transformants and all five independent growth hormonereceptor cell lines isolated showed the same effect. A similar resultwas obtained with a second set of independently isolated rabbit growthhormone receptor transfectants. Rowlinson et al. (1995) observed asimilar effect, suggesting that IL-3/GH-independent survival is aconsequence of the transformation procedure. Although the growth hormonereceptor cell lines did not require IL-3 or hGH for growth, they stillshowed a robust proliferative response to IL-3 and hGH. The practicaleffect of the higher absorbance values in the absence of hGH is todecrease the “window” of the hGH response (the difference between themaximum and minimum absorbance values). This window consistently rangedfrom 40-70% of the zero growth factor values, similar to what wasreported by Rowlinson et al.(1995).

Pituitary GH and wild-type rhGH prepared by us had similar dose responsecurves in the bioassay with EC₅₀s ranging from 0.6-1.2 ng/ml indifferent experiments (Table 1). TABLE 1 Bioactivities of hGH and hGHCysteine Muteins Form Mean EC₅₀ EC₅₀ Range Assyed (ng/ml) (ng/ml)¹Pituitary hGH Monomer   1 +/− 0.1 (N = 11) 0.75-1.2  RhGH Monomer 0.8+/− 0.3 (N = 3) 0.6, 0.8, 1.1 T3C Dimer 1.4 +/− 0.6 (N = 7) 0.75-2.5 S144C Monomer 1.6 +/− 0.8 (N = 5) 1.1, 1.1, 1.5, 2.2, 2.7 T148C-lotAMonomer 0.5 (N = 2) 0.4, 0.5 T148C-lotB Monomer 2.5 +/− 1.0 (N = 4) 1.5,1.9, 3.1, 3.6N/A, not applicable¹EC₅₀ values from individual experiments. An EC₅₀ range is shown whenN > 5.

EXAMPLE 4 Construction of hGH Cysteine Muteins

The cysteine substitution mutation T135C was constructed as follows. Themutagenic reverse oligonucleotide BB28 (5>CTGCTTGAAGATCTGCCCACACCG GGGGCTGCCATC>3)(SEQ.ID.NO. 10) was designed to change the codon ACT forthreonine at amino acid residue 135 to a TGT codon encoding cysteine andto span the nearby Bgl II site. This oligo was used in PCR along withBB34 (5>GTAGCGCAGGCCTTCCCAACC ATT>3)(SEQ.ID.NO. 11) which anneals to thejunction region of the ompA-GH fusion gene and is not mutagenic. The PCRwas performed in a 50 μl reaction in 1×PCR buffer (Perkin-Elmer buffercontaining 1.5 mM MgCl₂), 200 μM concentration of each of the fournucleotides dA, dC, dG and dT, with each oligonucleotide primer presentat 0.5 μM, 5 pg of pBBT112 (described above) as template and 1.25 unitsof AmpliTaq DNA Polymerase (Perkin-Elmer) and 0.125 units of Pfu DNAPolymerase (Stratagene). Reactions were performed in a RobocyclerGradient 96 thermal cycler (Stratagene). The program used entailed: 95°C. for 3 minutes followed by 25 cycles of [95° C. for 60 seconds, 45° C.or 50° C. or 55° C. for 75 seconds, 72° C. for 60 seconds, followed by ahold at 60° C. The PCR reactions were analyzed by agarose gelelectrophoresis which showed that all three different annealingtemperatures gave significant product of the expected size; ˜430 bp. The45° C. reaction was “cleaned up” using the QIAquick PCR Purification Kit(Qiagen) and digested with Bgl II and Pst I. The resulting 278 bp BglII-Pst I fragment, which includes the putative T135C mutation, wasgel-purified and ligated into pBBT111, the pUC19 derivative carrying thestII-GH fusion gene (described above) which had been digested with BglII and Pst I and gel-purified. Transformants from this ligation wereinitially screened by digestion with Bgl II and Pst I and subsequentlyone clone was sequenced to confirm the presence of the T135C mutationand the absence of any additional mutations that could potentially beintroduced by the PCR reaction or by the synthetic oligonucleotides. Thesequenced clone was found to have the correct sequence throughout theBgl II-Pst I segment.

The substitution mutation S132C was constructed using the protocoldescribed above for T135C with the following differences: mutagenicreverse olignucleotide BB29(5>CTGCTTGAAGATCTGCCCAGTCCGGGGGCAGCCATCTTC>3)(SEQ.ID.NO. 12) was usedinstead of BB28 and the PCR reaction with annealing temperature of 50°C. was used for cloning. One of two clones sequenced was found to havethe correct sequence.

The substitution mutation T148C was constructed using an analogousprotocol but employing a different cloning strategy. The mutagenicforward oligonucleotide BB30(5>GGGCAGATCTTCAAGCAGACCTACAGCAAGTTCGACTGCAACTCACACAAC>3)(SEQ.ID.NO. 13)was used in PCR with the non-mutagenic reverse primer BB33(5>CGCGGTACCCGGGATCCGATTAGAATCCACAGCT>3)(SEQ.ID.NO. 14) which anneals tothe most 3′ end of the GH coding sequence and spans the Bam H1 siteimmediately downstream PCR was performed as described above with theexception that the annealing temperatures used were 46, 51 and 56° C.Following PCR and gel analysis as described above the 46 and 51° C.reactions were pooled for cloning. These were digested with Bam H1 andBgl II and the resulting 188 bp fragment was gel-purified and clonedinto pBBT111 which had been digested with Bam H1 and Bgl II, treatedwith calf alkaline phosphatase (Promega) according to the vendorprotocols, and gel-purified. Tranformants from this ligation wereanalyzed by digestion with Bam H1 and Bgl II to identify clones in whichthe 188 bp Bam H1-Bgl II mutagenic PCR fragment was cloned in the properorientation: because Bam H1 and Bgl II generate compatible ends, thiscloning step is not orientation specific. Five of six clones tested wereshown to be correctly oriented. One of these was sequenced and was shownto contain the desired T148C mutation. The sequence of the remainder ofthe 188 bp Bam H1-Bgl II mutagenic PCR fragment in this clone wasconfirmed as correct.

The construction of the substitution mutation S144C was identical to theconstruction of T148C with the following exceptions. Mutagenic forwardoligonucleotide BB31(5>GGGCAGATCTTCAAGCAGACCTACTGCAAGTTCGAC>3)(SEQ.ID.NO. 15) was usedinstead of BB30. Two of six clones tested were shown to be correctlyoriented. One of these was sequenced and was shown to contain thedesired S144C mutation. The sequence of the remainder of the 188 bp BamH1-Bgl II mutagenic PCR fragment in this clone was also confirmed ascorrect.

A mutation was also constructed that added a cysteine residue to thenatural carboxyterminus of GH. The construction of this mutation, termedstp 192C, was carried out using the procedures described above forconstruction of the T148C mutein but employed different oligonucleotideprimers. The mutagenic reverse oligonucleotide BB32(5>CGCGGTACCGGATCCTrAGCAGAAGCCACAGCTGCCCTCCAC>3)(SEQ.ID.NO. 16) whichinserts a TGC codon for cysteine between the codon for the carboxyterminal phe residue of GH and the TAA translational stop codon andspans the nearby Bam H1 site was used along with BB34 which is describedabove. Following PCR and gel analysis as described above the 46° C.reaction was used for cloning. Three of six clones tested were shown tobe correctly oriented. One of these was sequenced and was shown tocontain the desired stp192C mutation. The sequence of the remainder ofthe 188 bp Bam H1-Bgl II mutagenic PCR fragment in this clone wasconfirmed as correct.

The substitution mutation S100C was constructed using mutagenic reverseoligonucleotide BB25(5>GTCAGAGGCGCCGTACACCAGGCAGTTGGCGAAGAC>3)(SEQ.ID.NO. 17) which altersthe AGC codon for serine at amino acid residue 100 to a TGC codonencoding cysteine. BB25 also spans the nearby Nar I site. PCR reactionsusing BB25 and BB34 were carried out using the PCR protocol describedabove for the construction of the T135C mutation. Following gel analysisof the PCR products, the product of the 50° C. annealing reaction wascleaned up using the QLAquick PCR Purification Kit (Qiagen), withdigested with Pst I and Nar I. The resulting 178 bp fragment wasgel-purified and ligated into pBBT111 which had been digested with Pst Iand Nar I and gel-purified. Plasmids isolated from transformants fromthis ligation were screened by digestion with Pst I and Nar I andsubsequently one plasmid was sequenced to confirm the presence of theS100C mutation and the absence of any other mutations in the 178 bp Pst1-Nar I segment.

The substitution mutation A98C was constructed using the proceduredescribed above for S100C with the following differences: the mutagenicreverse oligonucleotide BB26(5>GTCAGAGGCGCCGTACACCAGGCTGTrGCAGAAGACACTCCT>3)(SEQ.ID.NO. 18) was usedfor PCR in place of BB25 and the PCR reaction performed with anannealing temperature of 45° C. was used for cloning. One clone wassequenced and found to have the correct sequence.

The substitution mutation A34C was constructed as follows. The mutagenicreverse oligo BB23(5>GCGCTGCAGGAATGAATACTTCTGTTCCTTTGGGATATAGCATTCTTCAAACTC>3)(SEQ.ID.NO.19) was designed to change the GCC codon for alanine at amino acidresidue 34 to a TGC codon encoding cysteine and to span the Pst I site.This oligonucleotide was used in PCR reactions along with BB11(5>CCCCCTCTAGACATATGAAGAAGAACATCGCATTCCTGCTGGCATCTATGTTCGTTTTCTCTATCG>3)(SEQ.ID.NO. 20)which anneals to the 5′ end of the coding sequence of the stII leadersequence and spans the Nde I site that overlaps the initiator methioninecodon.

PCR reactions were performed in 50 μl in 1×PCR buffer (Promega)containing 1.5 mM MgCl₂, 200 μM concentration of each of the fournucleotides dA, dC, dG and dT, with each oligonucleotide primer presentat 0.2 μM, 1 ng of pBBT111 (described above) as template and 0.8 unitsof Tac DNA Polymerase (Promega) and 0.33 units of Pfu DNA Polymerase(Stratagene). Reactions were performed in a Robocycler Gradient 96thermal cycler (Stratagene). The program used entailed: 96° C. for 3minutes followed by 25 cycles of [95° C. for 60 seconds, 50° C. or 55°C. or 60° C. for 75 seconds, 72° C. for 60 seconds] followed by a holdat 6° C. The PCR reactions were analyzed by agaraose gel electrophoresiswhich showed that all three different annealing temperatures gavesignificant product of the expected size; ˜220 bp. The 50 and 55° C.reactions were pooled, “cleaned up” using the QIAquick PCR PurificationKit (Qiagen) and digested with Nde I and Pst I. The resulting 207 bp NdeI-Pst I fragment, which includes the putative A34C mutation, wasgel-purified and ligated into pBBT111, which had been digested with NdeI and Pst I, treated with alkaline phosphatase, and gel-purified.Transformants from this ligation were initially screened by digestionwith Nde I and Pst I and subsequently one clone was sequenced to confirmthe presence of the A34C mutation and the absence of any additionalmutations that could potentially be introduced by the PCR reaction or bythe synthetic oligonucleotides. The sequenced clone was found to havethe correct sequence throughout the Nde I-Pst I segment

The substitution mutation S43C was constructed using the protocoldescribed above for A34C with the following differences: mutagenicreverse oligonucleotide BB24(5>GCGCTGCAGGAAGCAATACTTCTGTTCCTTTGG>3)(SEQ.ID.NO. 21) was used insteadof BB23. One clone was sequenced and shown to contain the correctsequence.

The substitution mutation T3C was constructed using two sequential PCRsteps. The first step created the desired mutation while the second stepextended the PCR product of the first reaction to encompass a usefulcloning site. The mutagenic forward oligonucleotide BB78(5>GCATCTATGTTCGTTTTCTCTATCGCTACCAACGCTTACGCATTCCCATGCATTCCCTTATCCAG>3)(SEQ.ID.NO. 22) was designed to changethe ACC codon for threonine at amino acid residue 3 to a TGC codonencoding cysteine and to span and anneal to the junction of the stII-hGHfusion gene. BB78 was used in the first step PCR along with BB33 whichis described above.

The first PCR reaction was performed in 50 μl in 1×PCR buffer (Promega)containing 1.5 mM MgCl₂, 200 μM concentration of each of the fournucleotides dA, dC, dG and dT, each oligonucleotide primer at 0.2 μM, 1ng of pBBT111 (described above) as template and 1.5 units of Tac DNAPolymerase (Promega) and 0.25 units of Pfu DNA Polymerase (Stratagene).Reactions were performed in a Robocycler Gradient 96 thermal cycler(Stratagene). The program used entailed: 96° C. for 3 minutes followedby 25 cycles of [94° C. for 60 seconds, 60° C. for 75 seconds, 72° C.for 60 seconds] followed by a hold at 6° C. The PCR reaction was ethanolprecipitated and the ˜630 bp product was gel-purified and recovered in20 μl 10 mM Tris-HCl (pH 8.5). An aliquot of this gel-purified fragmentwas diluted 100-fold and 2 μl of the diluted fragment was used astemplate in the second PCR step. The second PCR step employedoligonucleotides BB11 and BB33 (both described above) and used thereaction conditions employed in the first step PCR reaction. The secondstep PCR reaction was analyzed by agarose gel electrophoresis and theexpected ˜670 bp fragment was observed. The PCR reaction was “cleanedup” using the QIAquick PCR Purification Kit (Qiagen) and digested withNde I and Pst I. The resulting 207 bp Nde I-Pst I fragment, whichincludes the putative T3C mutation, was gel-purified and ligated intopBBT111, which had been digested with Nde I and Pst L treated withalkaline phosphatase, and gel-purified. Transformants from this ligationwere initially screened by digestion with Nde I and Pst I andsubsequently one clone was sequenced to confirm the presence of the T3Cmutation and the absence of any additional mutations that couldpotentially be introduced by the PCR reaction or by the syntheticoligonucleotides. The sequenced clone was found to have the correctsequence throughout the Nde I-Pst I segment.

The substitution mutation A105C was constructed using the technique of“mutagenesis by overlap extension” as described in general by Horton, R.M. pp251-261 in Methods in Molecular Biology, Vol. 15: PCR Protocols:Current Methods and Applications edited by White, B. A. (1993) HumanaPress, Inc., Totowa, N.J. With this technique two separate fragments areamplified from the target DNA segment as diagrammed in FIG. 1. Onefragment is produced with primers a and b to yield product AB. Thesecond primer pair, c and d, are used to produce product CD. Primers band c introduce the same sequence change into the right and left ends ofproducts AB and CD, respectively. Products AB and CD share a segment ofidentical (mutated) sequence, the “overlap”, which allows annealing ofthe top strand of AB to the bottom strand of CD, and the converse.Extension of these annealed overlaps by DNA polymerase in a subsequentPCR reaction using primers a and d with products AB and CD both added astemplate creates a full length mutant molecule AD.

With the exception of the use of different oligonucleotide primers, theinitial PCR reactions for the A105C construction were performedidentically to those described in the construction of T3C above. Theprimer pairs used were: (BB27+BB33) and (BB11+BB79). BB11 and BB33 aredescribed above. BB27 and BB79 are complementary mutagenicoligonucleotides that change the GCC codon for alanine at amino acidresidue 105 to a TGC codon encoding cysteine. The sequence of BB27 is(5>AGCCTGGTGTAC GGCTGCTCTGACAGCAACGTC>3)(SEQ.ID.NO. 23) and the sequenceof BB78 is 5>GACGTTGCTG TCAGAGCAGCCGTACACCAGGCT>3 (SEQ.ID.NO. 24). The(BB27×BB33) and (BB11×BB79) PCR reactions were ethanol precipitated,gel-purified, and recovered in. 20×10 mM Tris-HCl (pH 8.5). Thepreparative gel showed that the predominant product from each PCRreaction was of the expected size: ˜290 bp for the (BB27×BB33) reactionand ˜408 bp for the (BB11×BB79) reaction. These two mutagenizedfragments were then “spliced” together in a subsequent PCR reaction. Inthis reaction 1 μl of each of gel-purified fragments from the initialreactions were added as template and BB11 and BB33 were used as primers.Otherwise, the PCR reaction was performed using the same conditionsemployed for the initial (BB27×BB33) and (BB11×BB79) reactions. Analiquot of the secondary PCR reaction was analyzed by agarose gelelectrophoresis and the expected ˜670 bp band was observed. The bulk ofthe PCR reaction was then “cleaned up” using the QIAquick PCRPurification Kit (Qiagen) and digested with Bgl II and Pst I. Theresulting 276 bp Bgl II-Pst I fragment, which includes the putativeA105C mutation, was gel-purified and ligated into pBBT111, which hadbeen digested with Bgl II and Pst I, and gel-purified.

Transformants from this ligation were initially screened by digestionwith Bgl II and Pst I and subsequently one clone was sequenced toconfirm the presence of the A105C mutation and the absence of anyadditional mutations that could potentially be introduced by the PCRreaction or by the synthetic oligonucleotides. The sequenced clone wasfound to have the correct sequence throughout the Bgl II-Pst I segment.

For expression in E. coli as proteins secreted to the periplasmic space,the stII-hGH genes encoding the 11 muteins were excised from thepUC19-based pBBT111 derivatives as Nde I-Xba I fragments of ˜650 bp andsubcloned into the pCYB1 expression vector that had been used to expressrhGH. For expression experiments, these plasmids were introduced into E.coli W3110.

Using procedures similar to those described here and in Example 23, onecan construct other cysteine muteins of GR. The cysteine muteins can besubstitution mutations that substitute cysteine for a natural aminoresidue in the GH coding sequence, insertion mutations that insert acysteine residue between two naturally occurring amino acids in the GHcoding sequence, or addition mutations that add a cysteine residuepreceding the first amino acid, F1, of the GH coding sequence or add acysteine residue following the terminal amino acid residue, F191, of theGH coding sequence. The cysteine residues can be substituted for anyamino acid, or inserted between any two amino acids, anywhere in the GHcoding sequence. Preferred sites for substituting or inserting cysteineresidues in GH are in the region preceding Helix A, the A-B loop, theB-C loop, the C-D loop and the region distal to Helix D. Other preferredsites are the first or last three amino acids of the A, B, C and DHelices. Preferred residues in these regions for creating cysteinesubstitutions are F1, P2, P5, E33, K38, E39, Q40, Q46, N47, P48, Q49,T50, S51, S55, S57, T60, Q69, N72, N99, L101, V102, Y103, G104, S106,E129, D130, G131, P133, R134, G136, Q137, K140, Q141, T142, Y143, K145,D147, N149, S150, H151, N152, D153, S184, E186, G187, S188, and G190.Cysteine residues also can be inserted immediately preceding orfollowing these amino acids. One preferred site for adding a cysteineresidue would be preceding F1, which is referred to herein as *-1C.

One also can construct GH muteins containing a free cysteine bysubstituting another amino acid for one of the naturally ocurringcysteine residues in GH. The naturally occurring cysteine residue thatnormally forms a disulfide bond with the substituted cysteine residue isnow free. The non-essential cysteine residue can be replaced with any ofthe other 19 amino acids, but preferably with a serine or alanineresidue. A free cysteine residue also can be introduced into GH bychemical modification of a naturally occurring amino acid usingprocedures such as those described by Sytkowski et al. (1998),incorporated herein by reference.

Using procedures similar to those described in Examples 1-15, one canexpress the proteins in prokaryotic cells such as E. coli, purify theproteins, PEGylate the proteins and measure their bioactivities in an invitro bioassay. The GH muteins also can be expressed in eukaryotic cellssuch as insect or mammalian cells using procedures similar to thosedescribed in Examples 16-19, or related procedures well known to thoseskilled in the art. If secretion is desired, the natural GH signalsequence, or another signal sequence, can be used to secrete the proteinfrom eukaryotic cells.

EXAMPLE 5 Cystine Addition Improves Recovery of Cysteine Mutein T3C

E. coli strain W3110 containing the T3C mutation was grown overnight in3 ml of LB containing 100 μg/ml ampicillin. The saturated overnightculture was diluted to 0.03 O.D. at A₆₀₀ in 300 ml of LB containing 100μg/ml ampicillin and incubated at 37° C. in 2 L shake flasks. When theculture O.D. reached 0.420, 1.5 ml of 100 mM IPTG was added for a finalconcentration of 0.5 mM to induce expression of the recombinant protein.The induced culture was incubated at 37° C. overnight (˜16 h).

The induced overnight culture reached an O.D of 3.6 at A₆₀₀ and wassplit into 2×135 ml volumes and centrifuged using a Sorval RC-5centrifuge with a GSA rotor at 8,000 rpm for 10 minutes at 4° C. Thesupernatants were discarded and the cell pellets subjected to osmoticshock treatment as follows. The cell pellets were resuspended toapproximately 49 O.D. in 10 ml of ice cold Buffer A [20% sucrose, 10 mMTris-HCl pH 7.5] or Buffer B [20% sucrose, 10 mM Tris-HCl pH 7.5, 5.5 mMcystine (pH adjusted to 7.5-8.0)]. The pellets were resuspended bytrituration and vortexing and 1 ml of 0.25 M EDTA pH 8.0 was added togive a final concentration of ˜25 mM Resuspended cells were incubated onice for 30 minutes and centrifuged in an SS-34 rotor at 8,500 rpm for 10minutes at 4° C. The supernatants were discarded and the pelletsresuspended by trituration and vortexing in 10 ml of ice cold Buffer C[5 mM Tris-HCl pH 7.5] or Buffer D [5 mM Tris-HCl pH 7.5, 5 mM cystine(pH adjusted to 7.5-8.0)] and incubated on ice for 30 minutes. Theresuspended cell pellet was centrifuged in an SS-34 rotor at 8,500 rpmfor 10 minutes at 4° C. and the resultant supernatant (solubleperiplasmic fraction) was recovered and stored at −80° C. SDS-PAGEanalysis of the various supernatants revealed that by incorporatingcystine into the osmotic shock procedure, the amount of monomeric hGHspecies recovered in the soluble periplasmic fraction can besignificantly improved. In the sample treated with cystine, we observedthat a majority of the hGH ran as a single band of about 22 kDa on anon-reduced SDS-PAGE gel and that band co-migrated with pituitary hGH.In the absence of cystine, a number of different molecular weightspecies were observed in the monomeric range. Larger molecular weightprotein aggregates are visible when the gel is developed using Westernblotting.

EXAMPLE 6 General Method for Expression and Purification of GH CysteineAdded Variants

A standard protocol for isolation of the hGH cyteine muteins is asfollows. Cultures of W3110 E. coli strains containing the stII-hGHmutein plasmids were grown to saturation in LB containing 100 μg/mlampicillin at 37° C. The overnight cultures were typically diluted to0.025-0.030 O.D. at A₆₀₀ in LB containing 100 μg/ml ampicillin and grownat 37° C. in shake flasks. When culture O.D.'s reached 0.300-0.500 atA₆₀₀, IPTG was added to a final concentration of 0.5 mM to induceexpression. The induced cultures were incubated at 37° C. overnightInduced overnight cultures were pelleted by centrifugation in a SorvalRC-5 centrifuge at 8,000-10,000×g for 10 minutes at 4° C. and theresultant pellets subjected to osmotic shock based on the procedures ofKoshland and Botstein (1980) or Hsiung et al., (1986) depending on thesize of the culture. For osmotic shock, cell pellets were resuspended at25-50 O.D. in 20% sucrose, 10 mM Tris-HCl pH7.5. In certain instances 5or 25 mM EDTA pH 8.0 and/or 5 mM cystine (pH adjusted to 7.5-8.0) wereincluded in the resuspension buffer. The resuspended pellets wereincubated on ice for 15-30 minutes and centrifuged as above. Thesupernatants were discarded and the pellets resuspended at 25-50 O.D. atA₆₀₀ in 5 or 10 mM Tris-HCl pH 7.5. In certain instances 5 mM EDTA pH8.0 and/or 5 mM cystine (pH adjusted to 7.5-8.0) were included in theresuspension buffer. The resuspended pellets were incubated on ice for30 minutes and centrifuged as above. The resulting supernatant containsthe soluble periplasmic components and the pellet is comprised ofinsoluble periplasmic, and cell associated components.

Addition of cystine to the osmotic shock buffer resulted insignificantly improved recoveries and stabilization of many of thecysteine muteins. In the presence of cystine the cysteine muteins werelargely monomeric, whereas in the absence of cystine, a mixture of hGHmonomers, dimers and higher order aggregates were observed when thesamples were analyzed by non-reducing SDS-PAGE and Western blots.Presumably the higher order aggregates are a consequence of the addedcysteine residues in the proteins since they were greatly reduced orabsent with wild-type rhGH. Addition of cystine to the osmotic shockbuffer largely solved this problem. We believe cystine reacts with thefree cysteine residue in the muteins to form a stable mixed disulfide,thus preventing disulfide shuffling and aggregation. We have also testeda second dithiol, cystamine and have seen a similar stabilizing effecton cysteine muteins of hGH. Presumably other dithiols such as oxidizedglutathione would also lead to improved recoveries of proteinscontaining a free cysteine.

Of the 11 hGH muteins analyzed, six expressed at levels sufficient forisolation and purification. Non-reduced SDS-PAGE analysis of the osmoticshock supernatants for the A34C, S43C, A98C, S100C and S132C muteinsshowed little or no protein present at the correct molecular weight.Addition of cystine did not discernibly improve recovery of theseproteins. The relatively low expression levels of these muteins makes itdifficult to observe improved yields. Preliminary analyses of whole celllysates indicated that these mutant proteins were expressed, butinsoluble. The T3C, A105C, T135C, S144C, T148C and stp192C muteinsshowed moderate to good expression levels. Five of these cysteinemuteins (T3C, T135C, S144C, T148C and stp192C) showed significantlyimproved recoveries when cystine was present in the osmotic shockbuffer. A105C was not tested in the absence of cystine. General proteinpurification protocols are described below.

WT-rhGH was purified by ion exchange and gel filtration chromatography.The cysteine muteins were purified by an assortment of chromatographicprocedures including ion exchange, hydrophobic interaction (HIC), metalchelation affinity chromatographies (IMAC), Size ExclusionChromatography (SEC) or a combination of these techniques. Generally,the hGH mutein was captured from the fresh osmotic shock supernatantusing a Q-Sepharose fast flow resin (Pharmacia) equilibrated in 20 mMTris-HCl, pH 8.0. The column was washed with 20 mM Tris-HCl, pH 8.0 andbound proteins eluted with a linear 10 volume increasing salt gradientfrom 0 to 250 mM NaCl in 20 mM Tris-HCl, pH 8.0. Fractions containingthe hGH muteins were identified by SDS-PAGE and Western blotting. Thesefractions were pooled and stored frozen. Alternative resins can be usedto capture hGH muteins from the osmotic shock mixture. These includeHIC, cation ion exchange resins or affinity resins.

For hydrophobic interaction chromatography (HIC), the Q column pool wasthawed to room temperature and NaCl added to a final concentration of 2M. The pool was loaded onto the Butyl-Sepharose fast flow resinpreviously equilibrated in 2 M NaCl, 20 mM sodium phosphate, pH 7.5. hGHmuteins were eluted from the resin using a reverse salt gradient form 2M to 0 M NaCl in 20 mM phosphate, pH 7.5. Fractions containing the hGHmuteins were identified by SDS-PAGE and Western blotting, and pooled.

In some instances, the EEC pool was subsequently loaded directly onto anickel chelating resin (Qiagen) equilibrated in 10 mM sodium phosphate,0.5 M NaCl, pH 7.5. Following a wash step, muteins were recovered usinga 0-30 mM imidizole gradient in 10 mM sodium phosphate, 0.5 M NaCl, pH7.5. hGH has a high affinity for nickel, presumably through the divalentmetal-binding site formed by H18, H21 and E174. As a result, hGH can beobtained in highly pure form using a metal chelation column (Masano etal., 1989). All of the muteins tested bound tightly to the nickel columnand eluted at similar imidazole concentrations (around 15 mM) aswild-type rhGH.

EXAMPLE 7 Purification of T3C

The soluble periplasmic fraction prepared in the presence of cystine(Example 5) was loaded onto a 1 ml HiTrap Q-Sepharose column PharmaciaBiotech, Uppsala, Sweden) equilibrated in 10 mM Tris-HCl pH 8.0 and thebound proteins were eluted with a 20 column volume linear gradient of0-250 mM NaCl. The column load and recovered fractions were analyzed by14% SDS-PAGE. T3C eluted at a salt concentration around 170-200 mM whichwas significantly later than WT-hGH. Surprisingly, T3C mutein waspresent in a monomeric form when loaded on the column, but was recoveredpredominantly as a stable disulfide linked homodimer following elutionfrom the Q column. When reduced, the T3C mutein co-migrated on aSDS-PAGE gel with wild type hGH. Dimer formation occurred during the Qcolumn purification step. Cystine was not present in the Q column bufferor the buffers used for the other column steps. The T3 C dimer remainedintact throughout any further concentration or purification procedures.The pool from the Q column was adjusted to a final NaCl concentration of0.5 M and loaded onto 1 ml Ni-agarose column (Qiagen) previouslyequilibrated in 10 mM sodium phosphate, 0.5 M NaCl, pH 7.5. Following awash step, highly purified T3C dimers were recovered using a 0-30 mMimidizole gradient in 10 mM sodium phosphate, 0.5 M NaCl, pH 7.5. T3Cdimers were active in the bioassay (Example 10 and Table 1).

The A105C, T135C and stp192C muteins also were recovered from the Qcolumn as disulfide-linked dimers. It appears that certain muteins arecapable of forming stable disulfide-linked dimers, presumably throughthe added cysteine residue, once cystine is removed. We believemonomeric forms of these proteins could be stabilized by includingcystine or other dithiol compounds and/or cysteine blocking agents inall the column buffers and/or maintaining the pH of the buffers below 7to prevent disulfide rearrangement.

EXAMPLE 8 Purification of T148C

E. coli strain W3110 expressing the T148C mutein was grown overnight in3 ml of LB containing 100 μg/ml ampicillin. The saturated overnightculture was diluted to 0.025 O.D. at A₆₀₀ in 500 ml of LB containing 100μg/ml ampicillin and divided into 2×250 ml volumes and incubated at 37°C. in 2 L shake flasks. When the culture O.D. reached approximately0.35, 1.25 ml of 100 mM IPTG was added to each flask for a finalconcentration of 0.5 mM to induce expression of the recombinant protein.The induced culture was incubated at 37° C. overnight (˜16 h).

The induced overnight cultures reached an O.D of approximately 3.5 atA₆₀₀ and were centrifuged using a Sorval RC-5 centrifuge with a GSArotor at 8,000 rpm for 10 minutes at 4° C. The supernatants werediscarded and the cell pellets subjected to osmotic shock treatment asfollows. The cell pellets were resuspended to approximately 43 O.D. inice cold 20% sucrose, 10 mM Tris-HCl pH 7.5, 5.0 mM EDTA pH 8.0, 5.0 mMcystine (pH adjusted to 7.5-8.0) by trituration and vortexing.Resuspended cells were incubated on ice for 15 minutes, centrifuged inSorval centrifuge with an SS-34 rotor at 8,500 rpm for 10 minutes at 4°C. The supernatants were discarded and the pellets resuspended toapproximately 43 O.D. in ice cold 1 mM Tris-HCl pH 7.5, 5 mM EDTA pH8.0, 5 mM cystine (pH adjusted to 7.5-8.0) by trituration and vortexing.Resuspended cells were incubated on ice for 30 minutes and centrifugedin an SS-34 rotor at 8,500 rpm for 10 minutes at 4° C. and the resultantsupernatant (soluble periplasmic fraction) was recovered and stored at−80° C.

A second culture of T148C was prepared in a similar manner with thefollowing exceptions: 1) the induced culture volume was 200 ml andreached an O.D. of approximately 4.0 at A₆₀₀ after overnight incubationat 37° C., and 2) The cell pellet resuspensions were done atapproximately 30 O.D.s at A₆₀₀.

The soluble periplasmic fraction from the above preparations werecombined and dialyzed against 1 L of 10 mM Tris-HCl pH 8.0 overnight at4° C. The dialysis retentate was loaded onto a 5 ml HiTrap Q-Sepharosecolumn (Pharmacia Biotech, Uppsala, Sweden) equilibrated in 10 mMTris-HCl pH 8.0 and the bound proteins were eluted with a 20 columnvolume linear gradient of 0-250 mM NaCl. Column fractions were analyzedby 14% SDS-PAGE (Novex, San Diego, Calif.). T148C mutein eluted as asharp peak at a salt concentration around 60-80 mM. The appropriatefractions were pooled and concentrated on Centricon 10 concentrators(Amicon, Inc., Beverly, Mass.). The concentrated pool was furtherpurified on a Superdex 200 HR 10/30 column (Pharmacia Biotech Inc.,Uppsala, Sweden.) equilibrated in 20 mM sodium phosphate pH 7.5, 150 mMNaCl. The column fractions were again analyzed by SDS-PAGE and the T148Cmutein containing fractions were pooled. We observed that the T148Cremained monomeric after recovery from the Q-Sepharose column and elutedfrom the Superdex column at a molecular weight similar to wild type hGH.T148C prepared by Q-Sepharose and size exclusion chromatography isreferred to as Lot A.

Alternatively the Q-pool can be loaded directly onto a nickel chelatingresin (Qiagen) equilibrated in 10 mM sodium phosphate, 0.5 M NaCl, pH7.5. Following a wash step, T148C was recovered using a 0-30 mMimidizole gradient in 10 mM sodium phosphate, 0.5 M NaCl, pH 7.5. T148Cprotein prepared by Q-Sepharose and nickel affinity chromatography isreferred to as Lot B.

An osmotic shock T148C supernatant was prepared in the absence ofcystine and purification over a Q-Sepharose column was attempted. Theidentical-protocol was followed as described above except cystine wasabsent from the osmotic shock buffers. The T148C protein product elutedfrom the Q column over a broad range of salt concentrations, recoverieswere lower and the protein preparation was substantially less pure thanthe cystine treated T148C sample.

EXAMPLE 9 Purification of S144C

E. coli strain W3110 containing the S144C mutation was grown overnightin 3 ml of LB containing 100 μg/ml ampicillin. The saturated overnightculture was diluted to 0.03 O.D.s at A₆₀₀ in 250 ml of LB containing 100μg/ml ampicillin and incubated at 37° C. in a 2 L shake flask. When theculture O.D. reached approximately 0.3, 1.25 ml of 100 mM IPTG was addedfor a final concentration of 0.5 mM to induce expression of therecombinant protein. The induced culture was incubated at 37° C.overnight (˜16 h).

The induced overnight culture reached an O.D of approximately 3.3 atA₆₀₀ and was centrifuged using a Sorval RC-5 centrifuge with a GSA rotorat 8,000 rpm for 10 minutes at 4° C. The supernatant was discarded andthe cell pellet subjected to osmotic shock treatment as follows. Thecell pellet was resuspended to approximately 33 O.D. in ice cold 20%sucrose, 10 mM Tris-HCl pH 7.5, 5.0 mM cystine (pH adjusted to 7.5-8.0)by trituration and vortexing and 0.25 M EDTA pH 8.0 was added to a finalconcentration of 25 mM. Resuspended cells were incubated on ice for 30minutes, centrifuged in the Sorval RC-5 centrifuge with an SS-34 rotorat 8,500 rpm for 10 minutes at 4° C. The supernatants were discarded andthe pellets resuspended to approximately 33 O.D.s in ice cold 5 mMTris-HCl pH 7.5, 5 mM cystine (pH adjusted to 7.5-8.0) by triturationand vortexing. Resuspended cells were incubated on ice for 30 minutesand centrifuged in an SS-34 rotor at 8,500 rpm for 10 minutes at 4° C.and the resultant supernatant (soluble periplasmic fraction) wasrecovered and stored at −80° C.

The soluble periplasmic fraction from the above preparation was loadedonto a 5 ml HiTrap Q-Sepharose column (Pharmacia Biotech, Uppsala,Sweden) equilibrated in 10 mM Tris-HCl pH 8.0 and the bound proteinswere eluted with a 20 column volume linear gradient of 0-250 mM NaCl.Column fractions were analyzed by 14% SDS-PAGE (Novex, San Diego,Calif.) and S144C mutein-containing fractions, 125-150 mM NaCl, werepooled and frozen. The S144C mutein was monomeric.

EXAMPLE 10 Bioactivities of hGH Cysteine Muteins

The biological activities of the purified T3C, S144C and T148C cysteinemuteins were measured using the cell proliferation assay described inExample 1. Protein concentrations were determined using a Bradfordassay. All three muteins were biologically active. All of the muteinsreached the same level of maximum stimulation as pituitary hGH, withinthe error of the assay. The mean EC₅₀s for the T3C, S144C and T148Cmuteins were similar to that of pituitary hGH and rhGH (Table 1). Twoindependent preparations of the T148C mutein and one preparation each ofthe S144C and T3C muteins (all prepared in the presence of cystine) havebeen assayed multiple times (Table 1). The partially purified A105C,T135C and stp192C muteins also have been assayed and are biologicallyactive.

EXAMPLE 11 General Methods for Pegylation and Purification of CysteineMuteins

GH muteins can be PEGylated using a variety of cysteine-reactivePEG-maleimide (or PEG-vinylsulfone) reagents that are commerciallyavailable. Generally, methods for PEGylating the proteins with thesereagents will be similar to those described in WO 9412219 (Cox andMcDermott) and WO 9422466 (Cox and Russell), both incorporated herein byreference, with minor modifications. The recombinant proteins aregenerally partially reduced with dithiothreitol (DTT),Tris(2-carboxyethyl) phosphine-HCl (TCEP) or some other reducing agentin order to achieve optimal PEGylation of the free cysteine. The freecysteine is relatively unreactive to cysteine-reactive PEGs unless thispartial reduction step is performed. The amount of reducing agentrequired to partially reduce each mutein can be determined empirically,using a range of reducing agent concentrations at different pHs andtemperatures. Reducing agent concentrations typically vary from 0.5equal molar to 10-fold excess. Preferred temperatures are 4° C. to 37°C. The pH can range from 6.5 to 9.0 but is preferrably 7.5 to 8.5. Theoptimum conditions will also vary depending on the reductant and time ofexposure. Under the proper conditions, the least stable disulfides(typically intermolecular disulfides and mixed disulfides) are disruptedfirst rather than the more thermodynamically stable native disulfides.Typically, a 5-10 fold molar excess of DTT for 30 minutes at roomtemperature is effective. Partial reduction can be detected by a slightshift in the elution profile of the protein from a reversed-phasecolumn. In the case of a dimeric hGH, a shift in molecular weight isvisible by SDS-PAGE analysis run under non-reducing condition. Care mustbe taken not to over-reduce the protein and expose additional cysteineresidues. Over-reduction can be detected by reversed phase-HPLC (theover-reduced protein will have a retention time similar to the fullyreduced and denatured protein) and by the appearance of GH moleculescontaining two PEGs following the PEGylation reaction (detectable by amolecular weight change on SDS-PAGE). Wild type GH can serve as acontrol since it should not PEGylate under conditions that do not reducethe native intramolecular disulfides. Excess reducing agent can beremoved prior to PEGylation by size exclusion chromatography or bydialysis. TCEP need not be removed before addition of the PEGylationreagent as it is does not contain a free thiol group. The partiallyreduced protein can be reacted with various concentrations ofPEG-maleimide or PEG-vinylsulfone (typically PEG: protein molar ratiosof 1:1, 5:1, 10:1 and 50:1) to determine the optimum ratio of the tworeagents. PEGylation of the protein can be monitored by a molecularweight shift for example, using SDS-PAGE. The lowest amount of PEG thatgives significant quantities of mono-pegylated product without givingdi-pegylated product is typically considered optimum (80% conversion tomono-pegylated product is generally considered good). Generally,mono-PEGylated protein can be purified from non-PEGylated protein andunreacted PEG by size-exclusion, ion exchange, affinity, reversed phase,or hydrophobic interaction chromatography. Other purification protocolssuch as 2-phase organic extraction or salt precipitation can be used.The purified PEGylated protein can be tested in the cell proliferationassay described in Example 1 to determine its specific activity.

Experiments can be performed to confirm that the PEG molecule isattached to the protein at the proper site. This can be accomplished bychemical or proteolytic digestion of the protein, purification of thePEGylated peptide (which will have a large molecular weight) by sizeexclusion, ion exchange or reversed phase chromatography, followed byamino acid sequencing. The PEG-coupled amino acid will appear as a blankin the amino acid sequencing run.

EXAMPLE 12 Preparation and Purification of PEG-T3C

A preliminary titration study was performed to determine appropriatereducing agent and PEG reagent concentrations and to avoidover-reduction of the protein. We monitored partial reduction of theprotein by conversion of dimer to non-reduced monomer species onnon-reduced SDS-PAGE. One p aliquots of purified T3C dimer wereincubated with increasing concentrations of TCEP for 60 minutes at roomtemperature. The reactions were immediately analyzed by non-reducingSDS-PAGE. The amount of TCEP that yielded significant amounts of monomerT3C without overreducing and denaturing the protein was used forsubsequent experiments. TCEP is a convenient reducing agent for smallscale experiments because it does not interfere with the PEGylationreaction; thus the protein:TCEP mixture did not have to be dialyzedprior to PEG addition. At a larger scale inexpensive reducing agentssuch as DTT are preferred for reducing proteins. Generally, the proteinis treated with a reducing agent for an optimal amount of time. The pHof the reaction is then adjusted to 6.5 or below to limit disulfiderearrangements. The reducing agent is removed by dialysis or liquidchromatography. The pH is then readjusted to greater than 6.5 orpreferably 7.5 to 8.5 and the PEG reagent is added.

The titration experiments at pH 7.5 indicated that a five-fold molarexcess of TCEP for 60 minutes at room temperature converted the majorityof the T3C dimer species into properly disulfide-bonded monomer withoutover reducing the protein. Control experiments indicated that, asexpected, the T3C dimer needed to be reduced with TCEP to be PEGylated.These reaction conditions were then scaled to 100 μg reaction scale. A10-fold molar excess of 5 kDa maleimide-PEG (Fluka) was added to theT3C:TCEP mixture after 10 minutes and the PEGylation reaction wasallowed to proceed for 60 minutes at room temperature. The sample wasloaded quickly onto a 1 ml Q-Sepharose column equilibrated in 20 mMTris-HCl, pH 8.0. The column was washed with 20 mM Tris-HCl, pH 8.0 andbound proteins eluted with a linear 10 volume increasing salt gradientfrom 0 to 250 mM NaCl in 20 mM Tris-HCl, pH 8.0. Fractions containingmono-PEGylated T3C (a single PEG molecule attached to the T3C monomer)were identified by SDS-PAGE and Western blotting. These fractions werepooled and stored frozen. The presence of the PEG moiety decreases theprotein's affinity for the resin, allowing the PEGylated protein to beseparated from the non-PEGylated protein.

EXAMPLE 13 PEGylation of S144C and Other Cysteine Muteins

A preliminary titration study was also performed for S144C to determineappropriate reducing agent, PEG reagent concentrations and to avoidover-reduction of the protein as described in Example 12 for T3C. Thelarger scale PEGylation was carried out at pH 7.5 at room temperaturefor 2 h using a 2-fold molar excess of TCEP and a 10-fold molar excessof 5 kDa maleimide-PEG. SDS-PAGE analysis of the reaction mixture showedsome species with two or more PEGs present. These were separated fromthe mono-PEGylated S144C using a Q-Sepharose column as described inExample 12. Separately we performed a PEGylation reaction using 5 kDavinylsulfone-PEG (Fluka) which resulted in monopegylated S144C underidentical reducing conditions.

We have also performed small scale PEGylation reactions on T148C,stp192C and T135C, all of which yielded mono-PEGylated protein with 5kDa maleimide-PEG and/or 5 kDa vinylsulfone-PEG.

EXAMPLE 14 Bioactivity of the PEG-T3C and PEG-S144C hGH Proteins

The biological activity of the PEG-T3C and PEG-S144C proteins weremeasured in the cell proliferation assay (Example 1). The PEG-T3Cprotein showed a similar dose-response curve as pituitary hGH andnon-PEGylated T3C protein and reached the same level of maximalstimulation. The mean EC₅₀ value for PEG-T3C was 1.6 ng/ml (0.07 nM)(values of 1.3, 1.5, 1.7, 1.8 ng/ml in four experiments). Bioactivity ofthe PEG-T3C protein is at least 100-fold greater than that of rhGH thathas been PEGylated using non-specific NHS-PEG reagents (EC₅₀ of 440ng/ml (20 nM) as described in Clark et al., (1996).

The mean EC₅₀ value for PEG-S144C was 43 ng/ml (2 nM) (40 and 45 ng/mlin two experiments). Bioactivity of the PEG-S144C protein isapproximately 10-fold greater than that of rhGH that has been PEGylatedusing non-specific NHS-PEG reagents (EC₅₀ of 440 ng/ml (20 nM); Clark etal., 1996).

EXAMPLE 15 Construction of Disulfide-Linked Trimers and Disulfide-LinkedHigher Order Multimers of hGH

Additional hGH variants having more than one “free” cysteine could beconstructed and used to create higher order disulfide-linked multimersof hGH. These multimers could include trimers, tetramers, pentamers,hexamers, septamers, octamers, and any higher order multimers. Forexample, an hGH variant having two “free” cysteine residues could beconstructed by using recombinant DNA technology to recombine in vitroDNA plasmid vectors carrying individual “free” cysteine mutations.Alternatively, mutagenesis of an hGH cysteine variant could be employedto add an additional “free” cysteine mutation. Further iterations ofeither of these two procedures could be used to construct hGH variantshaving three or more “free” cysteines.

An hGH variant having two free cysteine residues could be used togenerate hGH trimers and higher order multimers as follows. Such avariant would be expressed in E. coli and recovered as a monomer in thesupernatant of an osmotic shock lysate as disclosed in Examples 5-9herein. Subsequent processing steps could then be employed to inducedi-sulfide bond formation, e.g. Q Sepharose chromatography as describedin Examples 6-9 herein. Under such conditions some hGH variants havingone free cysteine, such as T3C and stp192C, are converted virtuallyquantitatively to disulfide-linked dimers. Under the same or similarconditions intermolecular disulfide formation by an hGH variant havingtwo free cysteines, e.g. a double mutant that combined T3C and stp192C,would result in a polymerization of hGH molecules and the chain lengthof such polymers would in principle be unlimited. The chain length couldbe limited and to some extent controlled by addition to thepolymerization reaction of hGH molecules having only one free cysteinesuch as the T3C variant and/or the stp192C variant Disulfide bondformation between the growing polymer and a molecule having only onefree cysteine will “cap” or prevent further extension of one of the twopolymerization sites in the nascent polymer. A subsequent reaction of asecond hGH molecule that has only one free cysteine with the otherpolymerization site of that nascent polymer terminates polymerizationand fixes the length of that polymeric molecule. The average polymerlength could be controlled by the stoichiometry of the reactants, i.e.the ratio of hGH molecules with two free cysteines to hGH molecules withone free cysteine. Average shorter polymers would be favored by lowerratios and average longer polymers would be favored by higher ratios.More complex “branched” polymers could be constructed from reactionsinvolving hGH variants with 3 or more free cysteines with hGH variantshaving only one free cysteine.

Discrete size classes of certain polymers could subsequently be purifiedby chromatographic methods such as size exclusion chromatography, ionexchange chromatography, hydrophobic interaction chromatography, and thelike. Similar procedures to those described for GH could be used tocreate disulfide-linked dimers and higher order multimers of EPO andalpha interferon.

EXAMPLE 16 Cloning, Expression and Purification of Baculovirus(BV)-Expressed Recombinant Human Erythropoietin (rEPO)

A. Cloning a cDNA encoding EPO. A cDNA encoding human EPO was cloned byPCR using forward primer BB45 (5>CCCGGATCCATGGGGGTGCACGAATGTCCTG>3)(SEQ.ID.NO. 25) and reverse primer BB47(5>CCCGA ATTCTATGCCCAGGTGGACACACCTG>3)(SEQ.ID.NO. 26). BB45 anneals tothe DNA sequence encoding the initiator methionine and amino terminalportion of the EPO signal sequence and contains a Bam HI site forcloning purposes (Jacobs et al., 1985; Lin et al., 1985). BB47 annealsto the 3′ untranslated region of the EPO mRNA immediately downstream ofthe translational stop signal and contains an Eco RI restriction sitefor cloning purposes. Total RNA isolated from the human liver cell lineHep3B was used in first strand synthesis of single-stranded cDNA forPCR.

For preparation of total cellular RNA, Hep3B cells (American TypeCulture Collection) were grown in Delbecco's Modified Eagle's media(DMEM) supplemented with 10% fetal bovine serum (FBS). EPO expressionwas induced by treating the cells for 18 h with 130 μM Deferoxamine or100 μM cobalt chloride. Both compounds have been shown to induce EPOmRNA and protein expression in Hep 3B cells (Wang and Semenza, 1993).RNA was isolated from the cells using an RNeasy Mini kit (Qiagen, Inc.),following the manufacturer's directions. Approximately 320 μg of totalRNA was isolated from 1.4×10⁷ cells treated with cobalt chloride and 270μg of total RNA isolated from 1.4×10⁷ cells treated with Deferoxamine.First strand synthesis of single-stranded cDNA was accomplished using a1st Strand cDNA Synthesis Kit for RT-PCR (AMV) from Boehringer MannheimCorp and random hexamers were used as the primer. Subsequent PCRreactions using the products of the first strand syntheses as templateswere carried out with primers BB45 and BB47. The expected ˜600 bp PCRproduct was observed when reaction products were run out on an agarosegel. Both RNA preparations yielded an EPO PCR product. The PCR productwas digested with Bam HI and Eco RI and cloned into vector pUC19 thathad been cut with Bam HI and Eco RI and treated with alkalinephosphatase. DNA sequencing identified a clone containing the correctcoding sequence for the EPO gene. This plasmid was designated pBBT131and used in the construction of EPO variants by site directedmutagenesis as described in Example 17.

B. Expression of BV rEPO in Insect Cells

For expression in insect cells the EPO cDNA in pBBT131 was modified atboth the 5′ and 3′ ends. At the 5′ end, the sequence CAAA was addedimmediately upstream of the initiator ATG to enhance translation. Thissequence comprises a proposed consensus translational initiationsequence for baculovirus (Ayres et al., 1994; Ranjan and Hasnain, 1995).At the 3′ end, DNA encoding the 8 amino acid FLAG epitope sequence(asp-tyr-lys-asp-asp-asp-asp-lys)(SEQ.ID.NO. 27) was added to provide apurification system. The FLAG epitope was fused to the EPO gene via aflexible linker: ser-gly-gly-ser-gly-gly-ser (SEQ.ID.NO. 28). Thesemodifications were made via PCR using oligonucleotide primers thatincorporated the desired additions to the EPO sequence. Oligonucleotideprimer BB63 (5>CGCGGATCCAAAATGGGGGTGCAC GAATGTCCT>3)(SEQ.ID.NO. 29) wasused to modify the 5′ end of the gene. BB63 adds the CAAA sequenceupstream of the ATG, anneals to the DNA sequence encoding the initiatormethionine and amino terminal portion of the EPO signal sequence, andcontains a Bam HI site for cloning purposes. The linker and FLAGsequences were added in two sequential PCR reactions using reverseprimers BB60 (5>GTCTTTGTAGTCCGAGCCTCCGCTTCCGCCCGATCT GTCCCCTGTCCTGCA>3)(SEQ.ID.NO. 30) and BB61(5>CGCGAATTCTTAMATCGTCATCGTCTTTGTAGTCCGAGCCTCC>3)(SEQ.ID.NO. 31). BB60anneals to 3′ of the EPO coding sequence and contains the fused peptidelinker sequence and a portion of the FLAG sequence. BB61 overlaps asegment of the BB60 sequence annealing to the junction of thelinker-FLAG sequence and adds the remainder of the FLAG sequencefollowed by a translational stop codon (TAA) and an Eco RI site forcloning purposes. The modified EPO cDNA was cloned as a Bam H-Eco RIfragment into pUC19 and the DNA sequence of this construct wasconfirmed. The resulting plasmid was designated pBBT132 and used in theconstruction of EPO variants as described in Example 17. For expressionin baculovirus, the “FLAG-tagged” EPO cDNA was excised from pBBT132 as˜630 bp Bam HI-Eco RI fragment, gel purified, and cloned into thebaculovirus transfer vector pBlueBac4.5 (Invitrogen) which had been cutwith Bam HI and Eco RI and treated with alkaline phosphatase. One clonewas picked for further use and designated pBBT138.

pBBT138 DNA was used to cotransfect Spodoptera frugiperda derived cellline Sf 9 along with linearized (Bsu36 I digested) Bac-N-Blue™(Invitrogen Corporation) baculovirus DNA. The Bac-N-Blue™ genome isengineered so that formation of plaque-forming viral particles requiresrecombination between the linear Bac-N-Blue™ DNA and the pBlueBac4.5vector, resulting in the incorporation of the cloned EPO gene into thebaculovirus genome. This obligate recombination also results inincorporation of a functional β-galactosidase gene into the baculovirusgenome. The co-transfection was performed according to the Invitrogen“Bac-N-Blue™ Transfection Kit” protocols using 2×10⁶ Sf 9 cells togenerate a ˜1 ml supernatant. Dilutions of this supernatant were assayedon Sf 9 cells at 27° C. for blue-plaque formation. Ten blue plaques werepicked and subcultured. Each plaque was used to inoculate 2.5×10⁶ Sf 9cells in a T25 flask containing 5 ml of Grace's Insect Mediasupplemented with 10% FBS. After 5 days the supernatants from theseinfected cells (the “P1” stocks) were collected and assayed by WesternBlot for EPO expression. The ten resultant supernatants were prepared inSDS sample buffer with the addition of 1% β mercaptoethanol (BME) andelectrophoresed on precast 14% Tris-glycine polyacrylamide gels (Novex).Uninfected SF9 cell supernatant was, included as a negative control.Following electrophoresis, the proteins were transferred onto 0.45 μmnitrocellulose (Novex). The nitrocellulose membrane was blocked in TrisBuffered Saline (TBS) with 0.05% Tween 20 and 4% powered milk (blockingbuffer). Anti-FLAG M2 mouse IgG₁ monoclonal antibody (Eastman KODAK) wasused at 1:1500 or 1:2500 dilution in blocking buffer and the blotroutinely incubated overnight at 4° C. Alkaline phosphatase conjugatedSheep Anti-Mouse IgG₁ (The Binding Site Limited) was diluted 1:1000 inblocking buffer and the blot incubated for 1 hour at room temperature.The Western blot was developed using NBT\BCIP color developmentsubstrate (Promega). Nine of the ten isolates were positive for EPOexpression. The molecular-weight of the BV rEPO protein wasapproximately 30 kDa under reducing conditions and consisted on onemajor and one to two minor bands in this molecular weight range, whichis consistent with a variably glycosylated protein.

Two of the positive supernatants were tested in the in vitro EPObioassay described in Example 16.D below. Both supernatants stimulatedproliferation of the EPO-dependent cell line in a dose-dependent manner,indicating that they contained active EPO. Control supernatants of mockinfected Sf 9 cells and the one baculovirus supernatant that wasnegative for EPO expression by Western blot showed no detectableactivity in the EPO bioassay.

In order to produce and purify larger amounts of wild type BV.rEPO, onepositive recombinant baculovirus, termed bvBBT138A, was chosen forfurther amplification. A 500 ml high titer viral stock was prepared bysubculturing the P1 stock of isolate 138A at 27° C. in a 500 ml spinnerflask culture of Sf 9 cells in Grace's Insect Media supplemented with10% FBS. Grace's Insect Media contains approximately 100 μM L-cystine.The supernatant from this culture was harvested after 7 days and foundto have a titer of ˜10⁸ plaque-forming-units/ml. An aliquot of thislysate, termed the “P2” stock, was subsequently used to infect a 500 mlculture for larger scale production of wild type BV rEPO. A 500 mlculture of Sf 9 cells in Grace's Insect Media supplemented with 10% FBSwas grown in a spinner flask to a titer of 1.0×10⁶/ml and then infectedwith bvBBT138A at a multiplicity of infection of 1. After 3 days thesupernatant from this culture was harvested and wild type BV rEPO-138Aprotein purified as described in Example 16.C.

C. Affinity Purification of Wild Type Baculovirus-Produced rEPO

The cell supernatant was clarified by centrifugation and 0.2 μMfiltration. Expression of wild type BV rEPO was confirmed by Westernblot analysis. Wild type BV rEPO was purified in a single step procedureusing Anti-FLAG M2 Affinity Gel (Eastman KODAK). Briefly, 5 ml of the M2affinity gel was washed with 5 column volumes of 50 mM Tris pH 7.4, 150mM NaCl (TBS), 5 column volumes of 0.1M glycine pH 3.5, thenequilibrated in TBS. The clarified baculoviral cell supernatant wasadjusted to 150 mM NaCl and the equilibrated resin was added. Batchloading was allowed to continue at 4° C. overnight on a roller bottleapparatus. After overnight incubation, the resin was recovered using aPharmacia XK 16/20 FPLC column and washed with TBS until the A280reached baseline. The bound protein was eluted with 0.1M glycine pH 3.5and fractions were collected and neutralized with 1.0M Tris pH 9.0.Column fractions were prepared in SDS-PAGE sample buffer with theaddition of 1% BME when desirable and electrophoresed on precast 14%Tris-glycine polyacrylamide gels. Fractions from the M2 affinity columnthat contained most of the BV rEPO were pooled and concentrated on aCentricon 10 spin concentrator (Amicon). The final yield of wild type BVrEPO, as determined using a Bradford protein assay kit (BIO-RADLaboratories, Inc.) and bovine serum albumin (BSA) as the standard, wasapproximately 360 μg. The protein was estimated to be greater than 90%pure by Coomassie Blue staining of SDS gels.

D. In Vitro Bioactivity of Wild Type Baculovirus rEPO

A cell proliferation assay using the human UT7/epo cell line (Komatsu etal., 1991) was developed to measure bioactivity of wild type BV rEPO.The human UV/epo cell line was obtained from Dr. F. Bunn of HarvardMedical School. This cell line is dependent upon EPO for cellproliferation and survival (Boissel et al., 1993). The cells weremaintained in Iscove's Modified Delbecco's Media (IMDM) supplementedwith 10% FBS, 50 units/ml penicillin, 50 μg/ml streptomycin and 1unit/ml rEPO(CHO (Chinese Hamster Ovary) cell-expressed; purchased fromR&D Systems, Inc.). For bioassays, the cells were washed three timeswith IMDM media and resuspended at a concentration of 1×10⁵ cells/ml inIMDM media containing 10% PBS, 50 units/ml penicillin and 50 μg/mlstreptomycin. Fifty μl (5×10³ cells) of the cell suspension werealiquotted per test well of a flat bottom 96 well tissue culture plate.Serial 3-fold dilutions of the protein samples to be tested wereprepared in phenol red-free IMDM media containing 10% FBS, 50 units/mlpenicillin and 50 μg/ml streptomycin. Fifty μl of the diluted proteinsamples were added to the test wells and the plates incubated at 37° C.in a humidified 5% CO₂ tissue culture incubator. Protein samples wereassayed in triplicate wells. After 60-72 h, 20 μl of CellTiter 96AQueous One Solution Reagent (Promega Corporation) was added to eachwell and the plates incubated at 37° C. in the tissue culture incubatorfor 14 h. Absorbance of the wells was read at 490 nm using a microplatereader. Control wells contained media but no cells. Mean absorbancevalues for the triplicate control wells were subtracted from mean valuesobtained for each set of triplicate test wells. Serial dilutions of CHOcell-expressed rEPO or wild type BV rEPO were analyzed in parallel.

The UT7/epo cell line shows a strong proliferative response to rEPO, asevidenced by a dose-dependent increase in cell number and absorbancevalues. Absorbance is proportional to cell number up to values of 2.0(Promega Corporation product specifications). In the absence of rEPO,the majority of UT7/epo cells die, giving absorbance values less than0.1. Commercial CHO cell-expressed rEPO and wild type BV rEPO preparedby us reached the same maximal level of stimulation, within the error ofthe assay, and had similar mean EC₅₀s in the bioassay of approximately0.4-0.5 ng/ml (table 2). EC₅₀ values for these proteins ranged from 0.21to 0.65 ng/ml in assays performed on different days (Table 2); thereforecomparisons between protein samples were made on samples analyzed on thesame day. These EC₅₀ values agree with values reported in the R&DSystems specifications for CHO rEPO (0.05-0.1 unit/ml or approximately0.4-0.8 ng/ml).

EXAMPLE 17 Construction, Expression, Purification and Bioactivity of EPOCysteine Muteins

A. Construction of EPO Cysteine Muteins.

Eight mutant EPO genes were constructed using site-directed PCR-basedmutagenesis procedures similar to those used to construct the GrowthHormone muteins described in Example 4 (Innis et al, 1990; Horton etal., 1993). We constructed four muteins at or near the two N-linkedglycosylation sites in the A-B loop (N24C, T26C, N38C and T40C), twomuteins at or near the N-linked glycosylation site in the B-C loop (N83Cand S85C), one mutein at the O-linked glycosylation site in phenolred-free IMDM media containing 10% FBS, 50 units/ml penicillin and 50μg/ml streptomycin. Fifty μl of the diluted protein samples were addedto the test wells and the plates incubated at 37° C. in a humidified 5%CO₂ tissue culture incubator. Protein samples were assayed in triplicatewells. After 60-72 h, 20 μl of CellTiter 96 AQueous One Solution Reagent(Promega Corporation) was added to each well and the plates incubated at37° C. in the tissue culture incubator for 1-4 h. Absorbance of thewells was read at 490 nm using a microplate reader. Control wellscontained media but no cells. Mean absorbance values for the triplicatecontrol wells were subtracted from mean values obtained for each set oftriplicate test wells. Serial dilutions of CHO cell-expressed rEPO orwild type BV rEPO were analyzed in parallel.

The UT7/epo cell line shows a strong proliferative response to rEPO, asevidenced by a dose-dependent increase in cell number and absorbancevalues. Absorbance is proportional to cell number up to values of 2.0(Promega Corporation product specifications). In the absence of rEPO,the majority of UT7/epo cells die, giving absorbance values less than0.1. Commercial CHO cell-expressed rEPO and wild type BV rEPO preparedby us reached the same maximal level of stimulation, within the error ofthe assay, and had similar mean EC₅₀ s in the bioassay of approximately0.4-0.5 ng/ml (Table 2). EC₅₀ values for these proteins ranged from 0.21to 0.65 ng/ml in assays performed on different days (Table 2); thereforecomparisons between protein samples were made on samples analyzed on thesame day. These EC₅₀ values agree with values reported in the R&DSystems specifications for CHO rEPO (0.05-0.1 unit/ml or approximately0.4-0.8 ng/ml).

EXAMPLE 17 Construction, Expression, Purification and Bioactivity of EPOCysteine Muteins

A. Construction of EPO Cysteine Muteins. Eight mutant EPO genes wereconstructed using site-directed PCR-based mutagenesis procedures similarto those used to construct the Growth Hormone muteins described inExample 4 (Innis et al, 1990; Horton et al., 1993). We constructed fourmuteins at or near the two N-linked glycosylation sites in the A-B loop(N24C, T26C, N38C and T40C), two muteins at or near the N-linkedglycosylation site in the B-C loop (N83C and S85C), one mutein at theO-linked glycosylation site in the C-D loop (S126C) and one mutein(*167C), that adds a cysteine between the natural carboxyterminal R166residue and the 15 amino acid carboxyterminal extension consisting ofthe peptide linker and FLAG sequences. The template for the mutagenicPCR reactions was plasmid pBBT131, in which the unmodified EPO gene iscloned as an Bam HI-Eco RI fragment into pUC19. PCR products weredigested with appropriate restriction endonucleases, gel-purified andligated with pBBT132 vector DNA that had been cut with appropriaterestriction enzymes, alkaline phosphatase treated, and gel-purified. Asdetailed above, pBBT132 is a pUC19 derivative carrying the clonedmodified (FLAG tagged) EPO gene. Transformants from these ligations weregrown up and plasmid DNAs isolated and sequenced. The sequence of theentire cloned mutagenized PCR fragment was determined to verify thepresence of the mutation of interest, and the absence of any additional

The substitution mutation N38C was constructed using the technique of“mutagenesis by overlap extension” as described in Example 4. With theexception of the use of different oligonucleotide primers, the initial,or “primary” PCR reactions for the N38C construction were performedidentically to those described in the construction of N24C above. Theprimer pairs used were (BB66+BB47) and (BB67+BB45). BB47 is describedabove. The forward, non-mutagenic, primer BB45(5>CCCGGATCCATGGGGGTGCACGAATGTCCTG>3)(SEQ.ID.NO. 35) anneals to the EPOsequence encoding the first seven amino acids of EPO. BB66 and BB67 arecomplementary mutagenic oligonucleotides that change the AAT codon forN38 to a TGT codon for cysteine. The sequence of BB66 is(5>AGCTTGAATGAGTGTATCACTGTCCCAGACACC>3)(SEQ.ID.NO. 36) and the sequenceof BB67 is (5>GGTGTCTGGGACAGTGATACACTCATTCAAGCT>3)(SEQ.ID.NO. 37). The(BB66×BB47) and (BB67×BB45) PCR reactions gave products of the expectedsizes: 420 bp for (BB66×BB47) and 220 bp for (BB67×BB45). The PCRproducts were ethanol-precipitated, gel-purified and recovered in 20 μl10 mM Tris-HCl as detailed above. These two mutagenized fragments werethen “spliced” together in the subsequent, or “secondary” PCR reaction.In this reaction 1 μl of each of the gel-purified PCR products of theprimary reactions were used as template and BB45 and BB47 were used asprimers. Otherwise, the reaction conditions identical to those used inthe primary reactions. An aliquot of the secondary PCR was analyzed byagarose gel electrophoresis and the expected band of ˜630 bp wasobserved. The bulk of the secondary PCR reaction was “cleaned up” usingthe QIAquick PCR Purification (Qiagen) according to the vendor protocol,digested with Sty I and Bsr GI (New England BioLabs) according to thevendor protocols, ethanol-precipitated, resuspended in 20 μl of 10 mMTris-HCl pH 8.5 and run out on a 2% agarose gel. The ˜400 bp Sty I-BsrGI fragment of interest was gel purified using a QIAEX II Gel ExtractionKit (Qiagen) according to the vendor protocol. This fragment was ligatedwith pBBT132 (described in Example 16) that had been cut with Sty I andBsr GI, treated with calf intestinal alkaline phosphatase (New EnglandBioLabs) and gel purified. The ligation reaction was used to transformE. coli and plasmids from resulting transformants were sequenced toidentify a clone containing the N38C mutation and having the correctsequence throughout the ˜400 bp Sty I-Bsr GI segment.

The substitution mutation T40C was constructed and sequence verifiedusing the procedures described above for N38C except that complementarymutagenic primers BB68 (5>AGCTTGAATGAGAATATCTGTGTCCCAGACACC>3)(SEQ.ID.NO. 38) and BB69 (5>GGTGTCTGGGACACAGATATTCTCATTCAAGCT>3)(SEQ.ID.NO. 39), which change the ACT codon for T40 to a TGTcodon for cysteine, replaced BB66 and BB67 respectively in the primaryPCR reactions.

The substitution mutation N83C was constructed and sequence verifiedusing the procedures described above for N38C except that complementarymutagenic primers BB70(5>GCCCTGTTGGTCTGCTCTTCCCAGCCGTGGGAGCCCCTG>3)(SEQ.ID.NO. 40) and BB71(5>CAGGGGCTCCCACGGCTGG GAAGAGCAGACCA ACAGGGC>3)(SEQ.ID.NO. 41), whichchange the AAC codon for N83 to a TGC codon for cysteine, replaced BB66and BB67, respectively, in the primary PCR reactions. The sizes of theproducts of the primary PCR reactions were also different. The(BB70×BB47) reaction gave, as predicted, a product of ˜300 bp and the(BB71×BB45) reaction gave, as predicted, a product of ˜360 bp.

The substitution mutation S85C was constructed and sequence verifiedusing the procedures described above for N38C except that complementarymutagenic primers primers BB72 (5>GCC CTGTTGGTCAACTCTTGCCAGCCGTGGGAGCCCCRG>3)(SEQ.ID.NO. 42) and BB73 (5>CAGGGGCTCCCACGGCTGGCAAGAGTRGACCAACAGGGC>3)(SEQ.ID.NO. 43), which change the TCC codonfor S85 to a TGC codon for cysteine, replaced BB66 and BB67 respectivelyin the primary PCR reactions. The sizes of the products of the primaryPCR reactions were also different. The (BB72×BB47) reaction gave, aspredicted, a product of ˜300 bp and the (BB73×BB45) reaction gave, aspredicted, a product of ˜360 bp.

The substitution mutation S126C was constructed using the proceduresdescribed above for N38C except that complementary mutagenic primersBB74 (5>CCAGATGCGGCCTGTGCTGCTCCACTC>3)(SEQ.ID.NO. 44) and BB75(5>GAGTGGAGCAGCACAGGCCGCATCTGG>3)(SEQ.ID.NO. 45), which change the TCAcodon for S126 to a TGT codon for cysteine, replaced BB66 and BB67respectively in the primary PCR reactions. The sizes of the products ofthe primary PCR reactions were also different The (BB74×BB47) reactiongave, as predicted, a product of ˜175 bp and the (BB75×BB45) reactiongave, as predicted, a product of ˜480 bp.

A mutation was also constructed that added a cysteine following thecarboxyterminal amino acid of the EPO coding sequence. This mutant,termed *167C was constructed as follows. A PCR reaction was carried outunder the conditions described above for the construction of the N24Cmutant, but employing oligonucleotides BB45 (see above) and reversemutagenic oligonucleotide BB77 (5>TTTGTAGTCCGAG CCTCCGCTTCCGCCCGAACATCTGTCCCCTGTCCTGCA>3)(SEQ.ID.NO. 46) and using 2.5 units of TaqPolymerase and 0.5 units of Pfu Polymerase. BB77 anneals to the terminal21 residues of EPO coding sequence and adds a TGT codon for cysteinefollowing the AGA codon for R166, which is the terminal amino acid inthe EPO coding sequence. BB77 also adds sequences encoding the sevenamino acid linker -ser-gly-gly-ser-gly-gly-ser-(SEQ.ID.NO. 27), and aportion of the FLAG epitope sequence. The ˜630 bp product of this PCRreaction was gel purified and used as template in a subsequent PCRreaction employing the same reaction conditions but using primers BB47and BB61 (5>CGCGAATTCTTATTTATCGTCATCGTCTTTGTAGTCCGAGCCTCC>3)(SEQ.ID.NO.30), which adds the remainder of the FLAG epitopesequence followed by a TAA stop codon and an Eco RI cloning site. The˜675 bp product of this PCR reaction was “cleaned up” using the QIAquickPCR Purification (Qiagen) according to the vendor protocol, digestedwith Sty I and Eco RI (New England BioLabs) according to the vendorprotocols, ethanol-precipitated, resuspended in 20 μl of 10 mM Tris-HClpH 8.5 and run out on a 2% agarose gel. The ˜86 bp Eco RI-Bsr GIfragment of interest was gel purified using a QIAEX II Gel ExtractionKit (Qiagen) according to the vendor protocol. This fragment was ligatedwith pBBT132 (described in Example 16) that had been cut with Eco RI andBsr GI, treated with calf intestinal alkaline phosphatase (New EnglandBioLabs) and gel purified. The ligation reaction was used to transformE. coli and plasmids from resulting transformants were sequenced toidentify a clone containing the *167C mutation and having the correctsequence throughout the ˜86 bp Eco RI-Bsr GI segment.

For expression in baculovirus, the EPO genes encoding the 8 muteins wereexcised from the pUC19-based pBBT132 derivatives as Bam HI-Eco RIfragments of ˜630 bp and subcloned into the pBlueBac4.5 baculovirustransfer vector used to express-wild type EPO.

Using procedures similar to those described here, one can constructother cysteine muteins of EPO. The cysteine muteins can be substitutionmutations that substitute cysteine for a natural amino residue in theEPO coding sequence, insertion mutations that insert a cysteine residuebetween two naturally occurring amino acids in the EPO coding sequence,or addition mutations that add a cysteine residue preceding the fistamino acid, A1, of the EPO coding sequence or add a cysteine residuefollowing the terminal amino acid residue, R166, of the EPO codingsequence. The cysteine residues can be substituted for any amino acid,or inserted between any two amino acids, anywhere in the EPO codingsequence. Preferred sites for substituting or inserting cysteineresidues in EPO are in the region preceding Helix A, the A-B loop, theBE loop, the C-D loop and the region distal to Helix D. Other preferredsites are the first or last three amino acids of the A, B, C and DHelices. Preferred residues in these regions for creating cysteinesubstitutions are A1, P2, P3, R4, L5, D8, S9, 125, T27, G28, A30, E31,H32, S34, N36, 139, D43, T44, K45, N47, Y49, A50, K52, R53, M54, E55,G57, Q58, G77, Q78, A79, S84, Q86, W88, E89, T107, R110, A111, G113,A114, Q115, K116, E117, A118, S120, P121, P122, D123, A124, A125, A127,A128, R131, T132, K154, Y156, T157, G158, E159, A160, T163, G164, D165,R166. Cysteine residues also can be inserted immediately preceding orfollowing these amino acids. Another preferred site for adding acysteine residue would be preceding A1, which we refer to as *-1C.

One also can construct EPO muteins containing a free cysteine bysubstituting another amino acid for one of the naturally-occurringcysteine residues in EPO. The naturally-occurring cysteine residue thatnormally forms a disulfide bond with the substituted cysteine residue isnow free. The non-essential cysteine residue can be replaced with any ofthe other 19 amino acids, but preferably with a serine or alanineresidue. A free cysteine residue also can be introduced into EPO bychemical modification of a naturally occurring amino acid usingprocedures such as those described by Sytkowski et al. (1998).

Using procedures similar to those described in Examples 16-19, one canexpress the proteins in eukaryotic cells (e.g., insect cells ormammalian cells), purify the proteins, PEGylate the proteins and measuretheir bioactivities in an in vitro bioassay. The EPO muteins also can beexpressed in prokaryotic cells such as E. coli using procedures similarto those described in Examples 1-15, or related procedures well known tothose skilled in the art

B. Insect Cell Expression of EPO-Cys Muteins.

For expression experiments, the eight plasmids encoding mutant EPO geneswere used to cotransfect Sf 9 cells along with linearized Bac-N-Blue™DNA using the procedures described above for plasmid pBBT138, whichencodes wild type EPO. The transfection supernatants were assayed on Sf9 cells for blue-plaque formation. For each mutant, ten blue plaqueswere picked and subcultured as described in Example 16. Five of the tenresulting supernatants were screened by SDS-PAGE and Western blot todetect expression of the EPO muteins. Western blot analysis was carriedout using the procedure described in Example 16 for wild type EPO.Western results showed that at least 3 of the 5 supernatants screenedfrom each clone contained FLAG-tagged EPO protein. Western blot resultsalso revealed that plasmids encoding muteins that should preventglycosylation at one of the N-linked glycosylation sites (N24C, T26C,N38C, T40C, N83C and S85C) yielded proteins with molecular weightsapproximately 2,200-2,800 daltons smaller than wild type BV rEPO. Incontrast, the S126 mutein at the O-linked glycosylation site, and *167C(C-terminal cysteine addition) yielded EPO proteins that co-migratedwith wild type BV rEPO. These results are consistent with theobservation that insect cells typically perform N-linked glycosylationand that sugar groups attached to O-linked glycosylation sites aregenerally small and cause minimal increases in a protein's molecularweight Insect cells are reported to perform O-linked glycosylation(Davies, 1995).

In order to purify larger amounts of the EPO muteins one positiverecombinant baculovirus isolate that encoded each mutein was chosen forfurther amplification. A 500 ml high titer viral stock was prepared fromeach of the mutant P1 stocks by subculturing as described above for thewild type isolate. An aliquot of each of these P2 lysates wassubsequently used to infect a 500 ml culture for larger scale productionof each of the EPO muteins using the procedures described for wild typeEPO in Example 16. The 500 ml supernatants from baculovirus infectedcells were subsequently sterile filtered and stored at 4° C. rEPO-Cysmuteins were purified using an anti-FLAG M2 Affinity Gel column (EastmanKodak), using a different batch of resin for each mutein to ensure nocross-contamination. 5 ml of resin was first washed with 50 mM Tris, 150mM NaCl, pH 7.4 (TBS), followed by 0.1 M Glycine pH 3.0, and finallyre-equilibrated in TBS. Sodium chloride was added to the baculovirusculture filtrate to a final concentration of 0.15 M before the affinityresin was added. The resin was batch-loaded on a roller bottle apparatusovernight at 4° C. The next morning the resin was captured using an XK16Pharmacia column and washed with TBS until the A280 reached baseline.The rEPO-Cys muteins were eluted with 0.1 M Glycine, pH 3.0. Fractions(1.5 ml) were collected into tubes containing 20 μL of 1 M Tris Base, pH9.0 to neutralize the solution. Based on the chromatograms and SDS-PAGEanalyses, fractions containing predominantly pure rEPO-Cys proteins werepooled and frozen Protein recoveries from the 500 ml supernatantcultures varied from 75 to 800 μg, depending upon the mutein (Table 2).

The purified rEPO-Cys muteins migrated with apparent molecular weightsof 27-30 kDa under non-reducing conditions, consistent with the proteinsbeing monomeric. The apparent molecular weights of the rEPO-Cys muteinsfell into two classes, presumably due to glycosylation differences.rEPO-Cys proteins containing mutations at the O-glycosylation site(S126C) and at the C-terminus (*167C) migrated at a position similar towild type BV rEPO. Proteins containing mutations that should preventglycosylation of one of the N-linked glycosylation sites (N24C, T26C,N38C, T40C, N83C and S85C proteins) migrated with apparent molecularweights of 27-28 kDa, slightly smaller than wild type BV rEPO.Surprisingly, the S85C protein migrated as a doublet under both reducingand non-reducing conditions; the major band of the doublet was theexpected size for a protein lacking glycosylation at one of the N-linkedglycosylation sites, while the minor band was the size expected forfully glycosylated wild-type BV rEPO. All of the original recombinantBaculovirus plaques isolated for the S85C mutein yielded rEPO proteinsthat migrated as doublets, suggesting that the doublet is due to partialglycosylation at N83 (or another amino acid) rather than contaminationof S85C with wild type BV EPO or S126C or *167C proteins. Several of themuteins, e.g., S126C, *167C, had small amounts of a second protein thatmigrated with an apparent molecular weight of 45-55 kDa, depending uponthe mutein. This is the molecular weight expected for disulfide-linkedrEPO dimers. These bands reacted with the anti-FLAG antibody in Westernblots and were absent when the SDS gels were run under reducingconditions. These data indicate that the 45-55 kDa bands representdisulfide-linked rEPO-Cys dimers.

C. Bioactivities of EPO Cysteine Muteins.

Biological activities of the purified rEPO-Cys muteins were measured inthe UT1/epo cell proliferation assay described in Example 16. Proteinconcentrations were determined using a Bradford assay. Because of thelarge number of samples the muteins were divided into two groups foranalysis: muteins T26C, T40C, N83C and S126C comprised one group andmuteins N24C, N38C, S85C and *167C comprised the second group. Muteinswithin a group were analyzed on the same day. Wild type BV rEPO and CHOrEPO were analyzed in parallel on the same days to control for interdayvariability in the assays. All of the muteins stimulated proliferationof UT7/epo cells and had EC₅₀s that were within three-fold of the wildtype rEPO control proteins. Mean EC for the N24C, T26C, N83C, S85C, S126and *167C muteins were similar to the mean EC₅₀s for wild type BV rEPOand CHO rEPO, averaging 0.3-0.8 ng/ml (Table 2). Mean EC₅₀s for the N38Cand T40C proteins were approximately 1 ng/ml (Table 2).

Bill et al., (1995) reported expression of the EPO cysteine muteins,N24C, N38C and N83C, in E. coli. In contrast to our results, theyreported that bioactivities of these cysteine muteins were significantlyreduced relative to wild type EPO. Bioactivity of the N38C mutein wasreduced to less than 20% of wild type EPO bioactivity and bioactivitiesof the N24C and N83C muteins were reduced to less than 5% of wild typeEPO bioactivity. EC₅₀s were not reported. Bill et al. (1995) postulatedthat the reduced activities of the cysteine muteins were due to improperfolding due to incorrect formation of disulfide bridges resulting fromthe extra cysteine residues introduced into the proteins. They did notemploy cystine or other cysteine-blocking agents in the solutions usedto purify the cysteine muteins.

Based upon the results of Bill et al. (1995), it was surprising that ourpurified N24C, N38C and N83C muteins had biological activities equal to,or within 3-fold of, wild type EPO. The data indicate that our methodsfor expression and purification of the EPO cysteine muteins results inproteins with superior in vitro biological activities compared to themethods employed by Bill et al. (1995). TABLE 2 Properties of Human EPOCysteine Muteins Expression Protein Recovery Mutation Mean EC₅₀ PlasmidEPO Protein μg/500 ml Location (ng/ml) EC₅₀ Range¹ — rEPO (CHO) — — 0.500.29-0.65 (N = 6) pBBT138 rEPO (BV) 345 — 0.37 0.21-0.51 (N = 6) pBBT150N24C 75 A-B loop 0.76 0.58, 0.85, 0.85 pBBT151 T26C 805 A-B loop 0.270.21, 0.22, 0.39 pBBT152 N38C 85 A-B loop 1.05 0.65, 1.20, 1.30 pBBT161T40C 102 A-B loop 0.95 0.60, 0.75, 1.50 pBBT162 N83C 96 B-C loop 0.500.42, 0.49, 0.60 pBBT153 S85C 135 B-C loop 0.72 0.50, 0.80, 0.85 pBBT154S126C 220 C-D loop 0.31 0.25, 0.25, 0.42 pBBT155 *167C 129 C-terminus0.61 0.51, 0.65, 0.68¹Data from individual experiments. A range is shown when N > 3.

EXAMPLE 18 PEGylation of EPO Cysteine Muteins

A. Small-Scale PEGylation of EPO Cysteine Muteins

Several cysteine muteins and wild type EPO were tested for their abilityto be PEGylated with a 5 kDa cysteine-reactive PEG following treatmentwith the reducing agent TCEP (Tris(2-carboxyethyl) phosphine-HCl). Atitration study was performed with the T26C mutein to identityappropriate TCEP and PEG reagent concentrations for PEGylation, whileavoiding over-reduction of the protein. We monitored partial reductionof the protein by SDS-PAGE, using wild type BV rEPO as a control.Increasing concentrations of TCEP (0.5 M to 6.0 M excess) in thepresence of a 10-40-fold molar excess of either vinylsulfone ormaleimide 5 kDa PEG (Fluka) were tested. The reactions were analyzed bynon-reducing SDS-PAGE. Wild type BV rEPO was treated in parallel as acontrol to identify partial reduction conditions that yieldedsignificant amounts of monoPEGylated EPO-Cys muteins (a single PEGmolecule attached to the EPO-Cys mutein), but no PEGylation of wild typeBV rEPO. A 5-fold molar excess of TCEP and a 30-fold molar excess of 5kDa vinylsulfone-PEG yielded significant amounts of monoPEGylated T26Cprotein without PEGylation of wild type BV rEPO.

Based on our findings with the T26C mutein, the following conditionswere used to PEGylate several other cysteine muteins. Aliquots (1-2 μ)of purified BV rEPO and EPO-Cys muteins were incubated for 1 hr with a5× molar excess of TCEP and a 30× molar excess of 5 kDa vinylsulfone PEGat pH 8.0 at room temperature. The reactions were stopped by dilutioninto SDS sample buffer (without reducing reagent) and analyzed bySDS-PAGE. Four cysteine muteins (N24C, T26C, S126C and *167C) werereadily monoPEGylated under these conditions, -as evidenced by theappearance of a new protein band migrating at approximately 35 kDa. The35 kDa species is the size expected for mono-PEGylated EPO; nodiPEGylated species, which are expected to have molecular weights ofapproximately 40 kDa, were detected for any of the EPO-cys muteins underthese reaction conditions. Control experiments indicated that thecysteine muteins needed to be reduced with TCEP in order to bePEGylated. Wild type BV EPO did not PEGylate under identical partialreducing conditions. These data indicate that the PEG molecule isattached to the free cysteine introduced into the N24C, T26C, S126C and*167C EPO-Cys muteins.

B. Preparation of PEG-T26C and PEG-S126C for Bioactivity Measurements.

We PEGylated larger quantities of the T26C and S126C proteins so thatthe PEGylated proteins could be purified for bioactivity measurements.PEGylation reactions were scaled to include 75 μg of each protein andthe same molar ratios of TCEP and 5 kDa-PEG used in the smaller 1 μgreactions. After 1 hour, the PEGylation mixture was diluted 10× with 20mM Tris-HCl, pH 8.0, 20% glycerol (Buffer A) and loaded immediately ontoa 1 ml Q-Sepharose column equilibrated in Buffer A The column was washedwith equilibration buffer and bound proteins eluted with a linear 10volume increasing salt gradient from 0 to 150 mM NaCl in 20 mM Tris-HCl,pH 8.0, 20% glycerol. The presence of the PEG moiety decreased theprotein's affinity for the resin, allowing the PEGylated protein to beseparated from the non-PEGylated protein. Fractions containingmono-PEGylated protein were identified by SDS-PAGE, followed byCoomassie Blue staining. Fractions containing PEG-T26C but no visibleunderivatized protein, were stored frozen at −80° C. and subsequentlyused in bioassays. Similar procedures were used to obtain purified PEG-S126C. A Western blot was performed to assess the purity of the PEG-T26Cand PEG-S 126C preparations used in bioassays. The Western blot wasperformed as described in Example 16. The Western blot gave strongsignals at the sizes expected for PEG-T26C and PEG-S126C, but failed todetect any protein migrating at the positions expected for theunPEGylated S126C and T26C proteins. From these results, we concludethere is little (<5%) unPEGylated protein present in the purifiedPEGylated muteins.

C. Bioactivities of PEG-T26C and PEG-S126C Cysteine Muteins

The biological activities of the purified PEG-T26C and PEG-S 126Cproteins were measured in the UT7/epo cell proliferation assay describedin Example 16. Protein concentrations of the PEG-EPO-Cys muteins werequantitated using a human EPO ELISA kit (R & D Systems), following themanufacturer's suggested directions. Protein concentrations of thenon-PEGylated muteins and wild type BV rEPO also were quantitated byELISA. The ELISA assay was performed on all the proteins on the sameday. Serial three-fold dilutions of the protein samples were preparedand analyzed in the bioassay. Unused material from each serial dilutionwas stored frozen at 80° C. and later analyzed in the ELISA to determinean accurate protein concentration for the stating material. Several ofthe serial dilutions for each protein sample were analyzed to ensurethat the protein concentration in at least one test sample fell withinthe linear range of the standard ELISA curve, which is 0.0025-0.2Units/ml or approximately 0.02-1.6 ng/ml. At least two of the serialdilutions for each sample fell within this linear range.

Biological activities for the PEG-T26C and PEG-S 126C muteins weresimilar to those of the non-modified T26C and S126C proteins and wildtype BV, rEPO. Mean EC₅₀s for the PEG-T26C and PEG-S 126C muteins weresimilar to the mean EC₅₀ values determined for the non-PEGylated T26Cand S126C muteins and wild type BV rEPO, ranging from 0.48-0.82 ng/ml(Table 3). Biologically active, PEGylated EPO proteins have not beendescribed previously. TABLE 3 Bioactivities of PEG-Cys EPO Muteins MeanEC₅₀ EC₅₀ Range¹ EPO Protein (ng/ml) (ng/ml) BV EPO 0.82 0.55, 0.95,0.95 T26C 0.73 0.50, 0.85, 0.85 PEG-T26C 0.58 0.35, 0.65, 0.75 S126C0.74 0.65, 0.75, 0.82 PEG-S126C 0.48 0.38, 0.52, 0.55¹Data from three experiments.

EXAMPLE 19 Expression and Purification of EPO Cysteine Muteins inMammalian Cells

The EPO cysteine muteins also can be expressed in mammalian cells andpurified from the conditioned media. The EPO cysteine muteins can beexpressed by transient transfection of mammalian cells or byconstructing stably transformed mammalian cell lines expressing the EPOcysteine muteins. For therapeutic applications in humans, it ispreferable that the EPO cysteine muteins be expressed without the linkerand FLAG sequences. Monkey COS cells (available from the American Typeculture Collection) can be used for transient transfection experiments,using procedures well known to those of skill in the art. A number ofcommercial sources, e.g., GIBCO/BRL sell lipid transfection reagents andprovide detailed protocols that can be used to express the EPO cysteinemuteins by transient transfection. Wild type EPO is manufactured for usein humans using stably transformed Chinese hamster ovary (CHO) cellsexpressing the protein. Stably transfected CHO cell lines are widelyused for high-level expression of recombinant proteins (Geisse et al.1996; Trill et al. 1995). In CHO cells, high level expression ofchromosomally integrated heterologous genes can be achieved by geneamplification. Typically the gene of interest is linked to a marker genefor which amplification is selectable. A variety of genes which provideselections for amplification have been described (Kaufman 1990) butmurine dihydrofolate reductase (dhfr) is frequently employed.Amplification of this gene confers resistance to the folate analogmethotrexate (MTX) and the level of resistance increases with dhfr genecopy number (Alt et al., 1978). Utility of MTX selection is enhanced bythe availability of CHO cell lines that are deficient in dhfr (Urlauband Chasin, 1980).

One skilled in the art can construct expression vectors for the EPOcysteine muteins that incorporate the murine dhfr gene into thecommercially available pCDNA3.1 expression vector (Invitrogen), whichincludes the neomycin phosphotransferase (NPT) gene that confersresistance to G418. The murine dhfr expression vector pdhfr2.9 isavailable from the American Type Culture Collection (catalogue No.337165). This dhfr gene is selectable in dhfr CHO cell lines and can beamplified by standard selections for MIX resistance (Crouse et al 1983).The dhfr coding sequence can be excised from pdhfr2.9 as a ˜900 bp BglII fragment and cloned into the unique Bam HI site of the polylinker ofthe expression vector pREP4 (Invitrogen). This construct will positionthe gene downstream of the strong RSV promoter, which is known tofunction in CHO cells (Trill et al, 1995) and upstream of apolyadenylation site derived from SV40. This dhfr expression cassettecan then be excised from pREP4 as a Sal I fragment since Sal I sitesclosely flank the promoter and polyA addition site. Usingoligonucleotide linkers this Sal I fragment can be cloned into theunique Bgl II site of pCDNA3.1.

For expression in mammalian cells such as COS or CHO cells, the EPOcysteine mutein cDNAs can be modified by PCR-based mutagenesis. At the5′ end one can add a Kozak consensus sequence prior to the translationalinitiator ATG codon in order to enhance translation in mammalian cells.At the 3′ end, the linker and FLAG sequences can be removed and thenatural coding sequence and translation termination signal restored.These modified EPO cDNAs can then be cloned as Bam HI-Eco RI fragmentsinto the polylinker of pCDNA3.1 for transient transfection experimentsor into the polylinker of the the pCDNA3.1::dhfr vector for stableexpression in mammalian cells. For mammalian cell expression, the mutantEPO genes encoding the cysteine muteins would be modified at the 5′ endto incorporate a Kozak consensus sequence (GCCACC) immediately precedingthe translational initiiatior ATG codon in order to enhance translationin mammalian cells. At the 3′ end the linker and FLAG sequences would beremoved and the natural coding sequence restored. These modificationscould be accomplished by a variety of mutagensis techniques that arewell known to those skilled in the art. PCR-based mutagensis proceduresbased on those described in Examples 4 and 17 could be employed. PCRprimers BB302 (5>CGCGGATCCGCCACCATGGGGGTGCACGAATGTCCT>3)(SEQ.ID.NO.47)and BB303 (5>CGCGAATTCTCATCTGTCCCCTGTCCTGCAGCC>3)(SEQ.ID.NO. 48) couldbe used to PCR the mutated EPO genes cloned in pUC19 or pBlueBac4.5 astemplates. Forward primer BB302 anneals to the EPO gene sequenceencoding the first seven amino acids of the EPO secretion signal, adds aKozak consensus sequence, GCCACC, immediately preceding thetranslational initiator ATG codon, and includes a Bam HI site forcloning purposes. The reverse primer BB303 anneals to sequences encodingthe carboxyterminal seven amino acids of the EPO coding sequence, adds aTGA translational stop codon following the EPO coding sequence andincludes a Eco RI site for cloning purposes. The products of theindividual PCR reactions using these primers and any mutant EPO genetemplate that has a mutation between these two primers, e.g. amino acidsubstitutions at amino acids 1 through 159 of the mature EPO codingsequence, will produce PCR products that can be purified, digested withBam HI and Eco RI and cloned into a vector suitable for expression inmammalian cells such as pCDNA3.1(+)(Invitrogen). PCR primers 302 and 303also could be used to modify EPO cysteine muteins in which a cysteineresidue is added preceding the first amino acid, A1, of the mature EPOcoding sequence, but distal to the EPO signal sequence. Mutations in theEPO sequence that encode the carboxyterminal seven amino acids, or the*167C mutation described in Example 17, would require the use ofalternative reverse primers in place of BB303. These primers would beindividually designed to include the mutated cysteine codon, at least 21nucleotides at the 3′ end of the oligo that exactly match the templatetarget, and the Eco RI site for cloning purposes. For example thereverse primer BB304 (5>CGCGAATTCRCAACATCT GTCCCCTGTCCTGCAGCC>3)(SEQ.ID.NO. 49) and BB302 could be used in a PCR reaction withthe mutated EPO *167C gene cloned in pUC19 or pBlueBac4.5 as template togenerate a modified mutant EPO gene encoding the *167C mutein that wouldbe suitable for expression in mammalian cell expression systems.

Endotoxin free plasmid DNAs are preferred for transfecting mammaliancells such as COS or dhfr⁻ CHO cells. Dhfr⁻ CHO cell lines can beobtained from a number of sources such as Dr. L. Chasin at ColumbiaUniversity (CHO K1 DUKX B11) or from the American Type CultureCollection (CHO duk⁻, ATCC No. CRL-9096). The cells can be cultured inF12/DMEM medium supplemented with 10% FBS, glutamine, glycine,hypoxanthine, and thymidine (Lucas et al., 1996). Transfections can becarried out by electroporation or by using transfection reagents wellknown to those of skill in the art such as LipofectAMINE (Gibco BRL),using the vendor protocols and or those described in the literature(Kaufman, 1990). One can select for dhfr⁺ transfectants in F12/DMEMsupplemented with 7% dialyzed FCS and lacking glutamine, glycine,hypoxanthine, and thymidine (Lucas et al., 1996). Alternatively one canselect for G418 resistance (encoded by the NPT gene of pCDNA3.1) andsubsequently screen transfectants for the dhfr⁺ phenotype. Dhfr⁺ clonescan be expanded in selection medium and culture supernatants screenedfor EPO cysteine mutein production using commercially available EPOELISA kits (available from R & D Systems) or by Western blot usinganti-EPO antisera (available from R&D Systems). Clones expressing theEPO cysteine mutein can then be pooled and subjected to multiple roundsof selection for MTX resistance at increasing drug concentration asdescribed by Kaufman (1990). After each round of MIX selection,individual clones can be tested for EPO cysteine mutein production.These procedures are well described in the literature and have been usedto express a variety of heterologous protein in CHO cells (reviewed inKaufman, 1990).

Preferably, the media used to grow the CHO cells expressing the EPOcysteine muteins should contain cystine or another cysteine blockingagent The EPO cysteine muteins can be purified from the conditionedmedium of CHO cells using protocols similar to those described by Imaiet al. (1990). After removal of the CHO cells by centrifugation, the EPOcysteine muteins can be purified from the supernatant by columnchromatography using techniques well known to those of skill in the art.The column chromatography steps employed for the purification of the EPOcysteine muteins could include Blue Sepharose, hydroxyapatite, reversedphase, hydrophobic interaction, size exclusion and ion-exchangechromatography.

EXAMPLE 20 Cloning, Expression and Purification of IFN-α2

A. Cloning DNA sequences Encoding IFN-α2.

There are at least 25 distinct IFN-α genes which encode proteins thatshare 70% or greater amino acid identity (Blatt et al.,1996). Due to thehigh degree of DNA sequence homology between IFN-α species, the IFN-α2gene was cloned in two steps. First, the IFN-α2 gene was amplified byPCR from human genomic DNA using primers corresponding to uniquesequences upstream and downstream of the IFN-α2 gene. This PCR productwas cloned and sequenced to confirm that it encodes the IFN-α2 gene.Subsequently, the IFN-α2 coding sequence was modified by PCR andsubcloned for expression in E. coli and site-directed mutagenesis. DNAencoding IFN-α2 was amplified by PCR from human genomic DNA (CLONTECH).PCR reactions were carried out with BB93 (5>CGCGAATTCGGATATGTAAATAGATACACAGTG>3) (SEQ.ID.NO. 50) and BB94(5>CGCAAGCTTAAAAGATMTAAATCGTGTCATGGT>3) (SEQ.ID.NO.51). BB93 anneals togenomic sequences ˜300 bp upstream (i.e. 5′ to) of the IFN-α2 codingsequence and contains an Eco RI site for cloning purposes. BB94 annealsto genomic sequences ˜100 bp downstream (i.e. 3′ to) of the IFN-α2coding sequence and contains a Hind III site for cloning purposes. Theresulting ˜1 kb PCR product was digested with Eco RI and Hind III andcloned into similarly digested, and alkaline phosphatased,pCDNA3.1(+)(Invitrogen). A clone having the correct DNA sequence forIFN-α2 (Henco et al, 1985) was identified and designated pBBT160.

For cytoplasmic expression in E. coli the cloned IFN-α2 gene of pBBT160was modified by PCR to incorporate a methionine codon immediately priorto the first residue (C1) of the mature IFN-α2 protein. A TAA stop codonwas added following the carboxyterminal residue, E165. At the same time,Xba I and Sal I sites were added and a Bgl II site was eliminated inorder to provide convenient restriction sites for subsequentmutagenesis. In this reaction pBBT160 was used as template and amplifiedby primers BB995>CGCAAGCITCATATGTGTGATCTGCCTCAAACCCACAGCCTGGGTTCTAGAAGGACCTTGATGCTC>3)(SEQ.ID.NO. 52) and BB100(5>CGCGAATTCTTATTCCTTACTTCTTAAACTTCTTGCAAGTTTGTCGACAAAGAAAAGGATCTCATGAT>3)(SEQ.ID.NO. 53). BB99 anneals to the 5′ end of the coding sequence ofmature IFN-α2 and encodes an initiator methionine preceding the firstamino acid of mature IFN-α2 BB99 introduces an Xba I site ˜30 bpdownstream of the initiator ATG, but the amino acid sequence of theprotein is unaltered. A Hind III site and an Nde I site, which overlapsthe ATG, were included for cloning purposes. The reverse primer, BB100,anneals to the 3′ end of the coding sequence and adds a TAA stop codon.BB100 introduces a Sal I site ˜30 bp upstream of the TAA codon andeliminates a naturally occurring Bgl II site located ˜15 bp furtherupstream. As a result, the naturally occurring Bgl II site located ˜185bp downstream of the initiator ATG becomes a unique site. None of thesealterations changed the amino acid sequence. An Eco RI site was addedimmediately downstream of the stop codon for cloning purposes. The ˜520bp PCR product was digested with Hind m and Eco RI, gel purified andcloned into similarly digested plasmid pCDNA3.1(+). One clone wasdetermined to have the correct DNA sequence. This plasmid was designatedpBBT164. For cytoplasmic expression in E. coli, the ˜520 bp Nde I-Eco RIfragment of pBBT164 was cloned into similarly digested expressionvector, pCYB1, (New England Biolabs). The plasmid vector pCYB1 allowsgenes to be expressed as unfused proteins or as fusion proteins; thisconstruct was created so that the protein is expressed as an unfusedprotein. The resulting plasmid was termed pBBT170 and encodesmet-IFN-α2. The Nde I-Eco RI fragment of pBBT164 containing themet-IFN-α2 sequence also was subcloned into Nde I-Eco RI-digested pUC18to generate plasmid pBBT168.

IFN-α2 also can be expressed in an active form in E. coli by secretioninto the periplasmic space (Voss et al., 1994). Secreted IFN-α2 lacks anN-terminal methionine and has an amino acid sequence identical tonaturally occurring IFN-α2. In order to express a secreted form ofIFN-α2, the leader sequence of the E. coli heat-stable enterotoxin(STII) gene (Picken et al., 1983) was fused to the coding sequence formature IFN-α2 via PCR. Because of its length, the STII sequence wasadded in two sequential PCR reactions. The first reaction used forwardprimer BB101 (5>GCATCTATGTTCGTTTTCTCTATCGCTACCAACGCTIACGCATGTGATCTGCCTCAAACCCAC AGC>3) (SEQ.ID.NO. 54) and reverse primer BB100 withpBBT164 (described above) as template. The 3′ end (21 bp) of BB101anneals to the 5′ end of the coding sequence of mature IFN-α2. The 5′segment (39 nucleotides) of BB101 encodes a portion of the STII leaderpeptide. The ˜550 bp PCR product of this reaction was gel purified andused as template for the second PCR reaction. The second PCR reactionused reverse primer BB100 and forward primer BB11(5′-CCCCCTCTAGACATATGAAG AAGAACATCGCAITCCTGCTGGCAT CTATGTTCGTTTTCTCTATCG-3′) (SEQ.ID.NO. 7). BB11 adds the remainder of the ST leaderpeptide and contains an Nde I site overlapping the initiator ATG of theSTII leader. The 590 bp product of this reaction was digested with Nde Iand Xba I. The 100 bp Nde I-XbaI fragment containing the STII leadersequence and amino-terminal ˜30 bp of IFN-α2, was gel-purified andligated with pBBT168 [pUC18::met-IFN-α2] that had been digested with NdeI and Xba I, treated with alkaline phosphatase and gel purified. In thisstep the ˜30 bp Nde I-Xba I amino-terminal segment of the met-IFN-α2gene is replaced with the PCR derived ˜100 bp Nde I-Xba I amino-terminalsegment of the STII-IFN-α2 PCR product. The sequence of the resultingpUC18::STII-IFN-α2 construct was confirmed and that plasmid vasdesignated pBBT177. For expression in E. coli, pBBT177 was digested withNde I and Eco RI and the ˜570 bp fragment containing the STII-IFN-α2gene was gel-purified and cloned into the expression vector pCYB1 underthe control of the tac promoter. The resulting plasmid was designatedpBBT178.

B. Expression of rIFN-α2 in E. coli.

pBBT170, which encodes met-IFN-α2, and parental vector, pCYB1, weretransformed into E. coli JM109. Experiments with these strains resultedin expression of met-IFNα-2. Secreted IFN-α2 is preferable tocytoplasmic met-IFN-α2 in that secreted IFN-α2 has the same amino acidsequence as naturally occurring IFN-α2.

For expression of secreted rIFN-α2, pBBT178 [pCYB1::STII-IFN-α2] and theparental vector pCYB1 were transformed into E. coli W3110. The resultingstrains were designated BOB202: W3110(pBBT178) and BOB130: W3110(pCYB1).We performed a series of experiments testing growth and rIFN-α2expression of BOB202 in phosphate- or MES-buffered LB media at initialpHs ranging from 5.0 to 7.0. Saturated overnight cultures were dilutedto ˜0.025 O.D. at A₆₀₀ in buffered LB containing 100 μg/ml ampicillinand incubated at 37° C. in shake flasks. When culture O.D.s reached˜0.3-0.5, IPTG was added to a final concentration of 0.5 mM to induceexpression of rIFN-α2. For initial experiments, cultures were sampled at0, 1, 3, 5 and ˜16 h post-induction. Samples of induced and uninducedcultures were analyzed by SDS-PAGE on precast 14% Tris-glycinepolyacrylamide gels stained with Coomassie Blue. In addition to theexpected ˜19 kDa processed form of IFN-α2, we observed a higher thanexpected molecular weight (˜21.5 kDa) form of the protein. The highermolecular weight form could result from lack of proteolytic processingof the STII leader peptide; the molecular weight of the leader peptideis consistent with this hypothesis. Our results indicated that lower pHenhanced accumulation of the correctly sized rIFN-α2 band. We observedthat at or above pH 6.5 the 21.5 kDa form was predominant, while at orbelow pH 6.0 a band of ˜19 kDa, which comigrated with an E.coli-expressed rIFN-α2 standard (Endogen, Inc.), was the predominantform of the protein. At or below pH 6.0, the 19 kDa band accounted forat least 80% and probably greater than 90% of the IFN-α2 expressed bythe E. coli cells. Based on these findings, further expressionexperiments used LB medium buffered with 100 mM MES to a pH of 5.5. Vosset al. (1994) also reported secretion of IFN-α2 to the E. coli periplasmusing the STII leader sequence. They also observed that the proportionof rIFN-α2 present in the 21.5 kDa band was reduced, and the proportionmigrating at 19 kDa was increased when culture pH was maintained at 6.7as compared to 7.0. At pH 7.0, Voss et al. (1994) reported that 10-30%of the STII:IFN-α2 fusion protein was processed to yield the 19 kDamature IFN-α2 protein. The percentage of correctly processed 19 kDaIFN-α2 protein could be increased to 50-60% by growing the E. coli at pH6.7. However, even at this pH, a substantial amount (40-50%) of theSTII: IFN-α2 fusion protein remained unprocessed, reducing the yield ofcorrectly processed 19 kDa IFN-α2. Voss et al. (1994) suggested that pH6.7 was optimal for maximizing the amount of secreted rIFN-α2 migratingat 19 kDa. Voss et al. (1994) varied several growth parameters toattempt to increase the percentage of correctly processed 19 kDa IFN-α2protein to greater than 50-60%, but were unsuccessful. Our data indicatethat lowering the pH to below 6.5, and preferably to 5.5 to 6.0,maximizes the ratio of cleaved (19 kDa) to uncleaved (21.5 kDa) rIFN-α2product. At these lower pHs, the percent of correctly processed 19 kDaIFN-α2 is increased to at least 80% and probably 90-100% of the totalIFN-α2 synthesized by the cells.

Cultures expressing rIFN-α2 were subjected to osmotic shock based on theprocedure of Koshland and Botstein (1980). This procedure ruptures theE. coli outer membrane and releases the contents of the periplasm intothe surrounding medium. Subsequent centrifugation separates the solubleperiplasmic components (recovered in the supernatant) from cytoplasmic,insoluble periplasmic, and cell-associated components (recovered in thepellet). Approximately 25-50% of the 19 kDa rIFN-α2 synthesized byBOB202 was recovered in the supernatant None of the 21.5 kDa form ofrIFN-α2 was observed in the soluble periplasmic fraction.

C. Large-Scale Expression and Purification of rIFN-α2 in E. coli:

In order to purify the wild type rIFN-α2 protein, fresh saturatedovernight cultures of BOB202 were inoculated at ˜0.02 OD @ A₆₀₀ in LB100 mM MES (pH5.5) containing 100 μg/ml ampicillin. Typically, a 325 mlculture was grown in a 2 liter shake flask at 37° C. in a gyrotoryshaker water bath at ˜160-200 rpm. When cultures reached a density of˜0.3-0.4 OD, IPTG was added to a final concentration of 0.5 mM Theinduced cultures were then incubated for ˜16 h. Cultures were subjectedto osmotic shock based on the procedure of Hsuing et al. (1986). Thecells were pelleted by centrifugation and resuspended at ˜25 OD/ml inice cold 20% sucrose, 10 mM Tris-HCl (pH 8.0). Resuspended cells wereincubated on ice for 15 min and centrifuged at 9500×g for 10 min.Pellets were then resuspended in ice cold 10 mM Tris-HCl pH (8.0),incubated on ice for 15 min and centrifuged at 9500×g for 10 min. Theresulting supernatant (the osmotic shock lysate) was either processedimmediately or stored at −80° C.

rIFN-α2 was purified as follows. The pH of the supernatant from theosmotic shock was adjusted to 3, centrifuged to remove any precipitate,and loaded onto a 5 ml Pharmacia HiTrap S-Sepharose column equilibratedin 20 mM MES pH 5.0 (Buffer A). The bound proteins were eluted with alinear salt gradient from 0-100% Buffer B (500 mM NaCl, 20 mM MES, 10%ethylene glycol). Column fractions were analyzed by non-reducingSDS-PAGE. rIFN-α2 eluted at approximately 225-235 mM NaCl. Fractionsthat were enriched for rIFN-α2 were pooled and further fractionated on a1 mL Cu⁺⁺ IMAC (Immobilized Metal Affinity Chromatography) Hi Trapcolumn previously equilibrated in 40 mM sodium phosphate pH 6.0, 1 MNaCl, 0.1% Tween 20. rIFN-α2 was eluted with a reverse pH gradient from5.5 to 4.1 in 40 mM sodium phosphate, 1 M NaCl, 0.1% Tween 20. rIFN-α2eluted after the gradient reached 100% buffer B, when the pH of theeluate finally reached pH 4.1. Fractions from the Cu⁺⁺ IMAC column thatcontained purified, properly folded rIFN-α2 were pooled and stored asfrozen aliquots at −80° C. A minor rIFN-α2 variant was detectable insome of the earlier eluting fractions. This variant, which results fromincomplete disulfide formation and is biologically active, has beendescribed previously (Khan and Rai, 1990). Fractions containing thisvariant were not added to the final pool of purified rIFN-α2. The finalyield of rIFN-α2, as determined by absorbance at 280 nm and by Bradfordanalysis, was about 400 μg from 250 ml of culture. Reduced IFN-α2migrates with a slightly larger apparent molecular weight thannon-reduced IFN-α2 when analyzed by SDS-PAGE (Morehead et al., 1984).This apparent molecular weight change is due to the reduction of thenative disulfides in IFN-α2. Our rIFN-α2 comigrated with the commercialrIFN-α2 standard under both reducing and non-reducing conditions.

D. In Vitro Bioactivity of Wild Type rIFN-α2.

IFN-α bioactivity can be measured using in vitro antiviral assays orcell proliferation inhibition assays. We developed a cell growthinhibition assay to measure bioactivity of wild type rIFN-α2. The humanDaudi B cell line (American Type Culture Collection) is sensitive to thegrowth inhibiting properties of IFN-α and is routinely used to measurebioactivity of IFN-α (Horoszewicz et al., 1979; Evinger and Peska,1981). Daudi cells were maintained in RPMI 1640 media supplemented with10% FBS, 50 units/ml penicillin and 50 μg/ml streptomycin. Forbioassays, the cells were washed three times with RPMI 1640 media andresuspended at a concentration of 4×10⁵ cells/ml in RPMI 1640 mediacontaining 10% FBS, 50 units/ml penicillin and 50 μg/ml streptomycin.Fifty μl (2×10⁴ cells) of the cell suspension were aliquotted per testwell of a flat bottom 96 well tissue culture plate. Serial 3-folddilutions of the protein samples to be tested were prepared in RPMI 1640media containing 10% FBS, 50 units/ml penicillin and 50 μg/mlstreptomycin. Fifty μl of the diluted protein samples were added to thetest wells and the plates incubated at 37° C. in a humidified 5% CO₂tissue culture incubator. Protein samples were assayed in triplicatewells. After 4 days, 20 μl of CellTiter 96 AQueous One Solution Reagent(Promega Corporation) was added to each well and the plates incubated at37° C. in the tissue culture incubator for 1-4 h. Absorbance was read at490 nm using a microplate reader. Control wells contained media but nocells. Mean absorbance values for the triplicate control wells weresubtracted from mean values obtained for each set of triplicate testwells. Serial dilutions of E. coli-expressed rIFN-α2 (Endogen, Inc.)were analyzed in parallel. IC₅₀s (the concentration of protein requiredfor half maximal growth inhibition) were calculated for each sample andused to compare bioactivities of the proteins.

Proliferation of the Daudi cell line is strongly inhibited by rIFN-α2,as evidenced by a dose-dependent decree in absorbance values. CommercialrIFN-α2 (Endogen) and wild type rIFN-α2 prepared by us reached the samemaximal level of growth inhibition, within the error of the assay, andhad similar mean IC₅₀s of 13-16 pg/ml (Table 4). IC₅₀ values for theseproteins ranged from 7-29 pg/ml in assays performed on different days(Table 4); therefore comparisons between proteins were made on samplesanalyzed on the same day.

EXAMPLE 21 Construction, Expression, Purification and Bioactivity ofIFN-α2 Cysteine Muteins

A. Construction of IFN-α2 Cysteine Muteins.

Seventeen mutant IFN-α2 genes were constructed using site-directedPCR-based mutagenesis procedures similar to those described in Example4. We constructed one mutein in the amino-terminal region proximal tohelix A [Q5C]; six muteins in the A-B loop [N45C, Q46C, F47C, Q48C, A50Cand 43C44 (an insertion of a cysteine between residues 43 and 44); onemutein [D77C] in the short, two residue, BC loop; four muteins in the CDloop [Q101C, T106C, E107C, and T108C]; three muteins in thecarboxy-terminal region distal to the E helix [S163C, E165C, and *166C(the addition of a cysteine residue to the natural carboxy-terminus)].We also constructed muteins C1S and C98S, which eliminate the naturallyoccurring, but nonessential C1-C98 disulfide (Lydon et al., 1985;Morehead et al., 1994). The C1S substitution in the amino-terminalregion proximal to helix A generates a free cysteine in the C helix(C98), whereas the C98S substitution in the C helix generates a freecysteine in the region proximal to helix A (C1).

For mutagenesis, PCR primer oligonucleotides were designed toincorporate nucleotide changes that resulted in the incorporation of acysteine residue at the chosen position within the IFN-α2 codingsequence. Where feasible, the mutagenic oligo also was designed to spana nearby restriction site that could be used to clone the mutagenizedPCR fragment into an appropriate plasmid. When no useful restrictionsite was located sufficiently near the position of the mutation, thetechnique of “mutagenesis by overlap extension” was employed (Horton etal., 1993). The template used for the mutagenic PCR reactions wasplasmid pBBT177 (described in Example 20) in which the STII-IFN-α2 geneis cloned as an Nde 1-Eco RI fragment into pUC18. The PCR products weredigested with appropriate restriction endonucleases, gel-purified andligated with pBBT177 vector DNA that had been cut with those samerestriction enzymes, alkaline phosphatase treated, and gel-purified.Transformants from these ligations were grown up and plasmid DNAsisolated and sequenced. The sequence of the entire cloned mutagenizedPCR fragment was determined to verify the presence of the mutation ofinterest, and the absence of any additional mutations that potentiallycould be introduced by the PCR reaction or by the syntheticoligonucleotide primers.

The substitution mutation Q5C was constructed using the technique of“mutagenesis by overlap extension” as described in Example 4. Theinitial, or “primary” PCR reactions for the Q5C construction wereperformed in a 50, reaction volume in 1× Promega PCR buffer containing1.5 mM MgCl₂, each primer at 0.2 μM, each of dATP, dGTP, dTTP and dCTPat 200 μM, 1 ng of template plasmid pBBT177 (described in Example 20),1.5 units of Taq Polymerase (Promega), and 0.25 units of Pfu Polymerase(Stratagene). Reactions were performed in a Robocycler Gradient 96thermal cycler (Stratagene). The reaction program entailed: 96° C. for 3minutes, 25 cycles of [95° C. for 1 minute, 60° C. for 1.25 minutes, 72°C. for 1 minute] followed by a hold at 6° C. The primer pairs used were[BB125×BB130] and [BB126×BB129]. BB125 (5>CTATGCGGCATCAGAGCAGATA>3)(SEQ.ID.NO. 55) anneals to the pUC18 vector sequence˜20 bp upstream of the cloned IFN-α2 sequence. BB126(5>TGTGGAATTGTGAGCGGATAAC>3)(SEQ.ID.NO. 56) anneals to the pUC18 vectorsequence ˜40 bp downstream of the cloned IFN-α2 sequence. BB129 andBB130 are complementary mutagenic oligonucleotides that change the CAAcodon for Q5 to a TGT codon for cysteine. The sequence of BB129 is(5>TGTGATCTGCCTTGTACCCACAGCCTG>3)(SEQ.ID.NO. 57) and the sequence ofBB130 is (5>CAGGCTGT GGGTACAAGGCAGATCACA>3)(SEQ.ID.NO. 58). The(BB125×BB130] and [BB126×BB129] PCR reactions gave products of theexpected sizes: ˜140 bp for [BB125×BB130] and ˜560 bp for [BB126×BB129].The PCR products were “cleaned up” using the QIAquick PCR PurificationKit (Qiagen) according to the vendor protocol, run out on a 2% agarosegel, gel-purified using a QIAEX II Gel Extraction Kit (Qiagen) accordingto the vendor protocol and recovered in 20 μl 10 mM Tris-HCl (pH 8.5).These two mutagenized fragments were then “spliced” together in thesubsequent, or “secondary” PCR reaction. In this reaction 2 μl of eachof the gel-purified PCR products of the primary reactions were used astemplate and BB125 and BB126 were used as primers. The reaction volumewas 100 μl and 2.5 units of Taq Polymerase and 0.5 units of PfuPolymerase were employed. Otherwise, the reaction conditions wereidentical to those used in the primary reactions. An aliquot of thesecondary PCR was analyzed by agarose gel electrophoresis and theexpected band of ˜670 bp was observed. The bulk of the secondary PCRreaction was “cleaned up” using the QIAquick PCR Purification (Qiagen),digested with Nde I and Xba I (New England BioLabs) according to thevendor protocols, “cleaned up” using the QIAquick PCR Purification Kitand run out on a 2% agarose gel. The ˜100 bp Nde I-Xba I fragment ofinterest was gel purified using a QIAEX II Gel Extraction Kit (Qiagen)according to the vendor protocol. This fragment was ligated with pBBT177(described in Example 20) that had been cut with Nde I and Xba I,treated with calf intestinal alkaline phosphatase (New England BioLabs)and gel purified. The ligation reaction was used to transform E. coliand plasmids from resulting transformants were sequenced to identify aclone containing the Q5C mutation and having the correct sequencethroughout the ˜100 bp Nde I-Xba I segment.

The substitution mutation C1S was constructed and sequence verifiedusing the protocols detailed above for Q5C except that complementarymutagenic primers BB128 (5>AGGCAGATCAGATGCGTAAGC>3)(SEQ.D.NO. 59) andBB127 (5>GCTTACGCATCTGATCTGCCT>3)(SEQ.ID.NO. 60), which change the TGTcodon for C1 to a TCT codon for serine, replaced BB130 and BB129respectively in the primary PCR reactions. The [BB125×BB1128] and[BB126×BB127] PCR reactions gave products of the expected sizes: 120 bpfor [BB125×BB128] and 570 bp for [BB126×BB127].

The substitution mutation N45C was constructed and sequence verifiedusing the protocols detailed above for Q5C with the followingdifferences. Complementary mutagenic primers BB134 (5>CTTTTGGAACTGGCAGCCAAACTCCTC>3)(SEQ.ID.NO. 61) and BB133(5>GAGGAGTITGGCTGCCAGTTCCAAAAG>3)(SEQ.ID.NO. 62), which change the AACcodon for N45 to a TGC codon for cysteine, replaced BB130 and BB129respectively in the primary PCR reactions. The [BB125×BB134] and[BB126×BB133] PCR reactions gave products of the expected sizes: ˜255 bpfor [BB125×BB134] and ˜440 bp for [BB126×BB133]. The product of thesecondary PCR reaction was “cleaned up” using the QIAquick PCRPurification (Qiagen), digested with Bgl II and Xba I (New EnglandBioLabs) according to the vendor protocols, “cleaned up” using theQIAquick PCR Purification Kit and run out on a 2% agarose gel. The ˜155bp Bgl II-Xba I fragment of interest was gel purified using a QIAEX IIGel Extraction Kit (Qiagen) according to the vendor protocol. Thisfragment was ligated with pBBT177 that had been cut with Bgl II and XbaI, treated with calf intestinal alkaline phosphatase (New EnglandBioLabs) and gel purified. The ligation reaction was used to transformE. coli and plasmids from resulting transformants were sequenced toidentify a clone containing the N45C mutation and having the correctsequence throughout the ˜155 bp Bgl II-Xba I segment.

The substitution mutation F47C was constructed and sequence verifiedusing the protocols detailed above for N45C with the followingdifferences. Complementary mutagenic primers BB136 (5>TTCAGCCTTTTGGCACTGGTTGCCAAA>3)(SEQ.ID.NO. 63) and BB135(5>TTTGGCAACCAGTGCCAAAAGGCTGAA>3)(SEQ.ID.NO. 64), which change the TTCcodon for F47 to a TGC codon for cysteine, replaced BB134 and BB133respectively in the primary PCR reactions. The [BB125×BB136] and[BB126×BB135] PCR reactions gave products of the expected sizes: ˜260 bpfor [BB125×BB136] and ˜435 bp for [BB126×BB135].

The insertion mutation 43C44 was constructed and sequence verified usingthe protocols detailed above for N45C with the following differences.Complementary mutagenic primers BB132 (5>TTCAGCCTTGGCACTGGTTGCCAAA>3)(SEQ.ID.NO. 65) and BB131(5>MGGCAACCAGTGCCAAAAGGCTGAA>3)(SEQ.ID.NO. 66), which insert a TGC codonfor cysteine between the codons encoding amino acid residues 43 and 44,replaced BB134 and BB133 respectively in the primary PCR reactions. The[BB125×BB132] and [BB126×BB131] PCR reactions gave products of theexpected sizes: ˜250 bp for [BB125×BB132] and ˜445 bp for [BB126×BB131].

The substitution mutation Q46C was constructed and sequence verifiedusing the protocols detailed above for N45C with the followingdifferences. Complementary mutagenic primers BB154 (5>AGCCTTTTGGAAACAGTTGCCAAACTC>3)(SEQ.ID.NO. 67) and BB153(5>GAGTTTGGCAACTGTMTCCAAAAGGCT>3)(SEQ.ID.NO. 68), which change the CAGcodon for Q46 to a TGT codon for cysteine, replaced BB134 and BB133respectively in the primary PCR reactions. The primary reactions wereperformed in a Perkin-Elmer GeneAmp® PCR System 2400 thermal cycler. Thereaction program entailed: 95° C. for 5 minutes, 30 cycles of [95° C.for 30 seconds, 62° C. for 30 seconds, 72° C. for 1 minute] followed by72° C. for 7 minutes and a hold at 4° C. The [BB125×BB154] and[BB126×BB153] PCR reactions gave products of the expected sizes: ˜260 bpfor [BB125×BB154] and ˜440 bp for [BB126×BB153]. The secondary PCRreaction was also performed in a Perkin-Elmer GeneAmp® PCR System 2400thermal cycler. This reaction program entailed: 96° C. for 5 minutes, 25cycles of [95° C. for 30 seconds, 60° C. for 30 seconds, 72° C. for 1minute] and a hold at 4° C. Following digestion with Bgl II and Xba I,the products of this reaction were cleaned up using the QIAquick PCRPurification Kit but not gel purified prior to ligation.

The substitution mutation Q48C was constructed and sequence verifiedusing the protocols detailed above for Q46C with the followingdifferences. Complementary mutagenic primers BB156 (5>GGTTTCAGCCTTACAGAACTGGTTGCC>3)(SEQ.ID.NO. 69) and BB155(5>GGCAACCAGTFCTGTAAGGCTGAAACC>3)(SEQ.ID.NO. 70), which-change the CAAcodon for Q48 to a TGT codon for cysteine, replaced BB154 and BB153respectively in the primary PCR reactions. The [BB125×BB156] and[BB126×BB155] PCR reactions gave products of the expected sizes: ˜265 bpfor [BB125×BB156] and ˜435 bp for [BB126×BB155].

The substitution mutation A50C was constructed and sequence verifiedusing the protocols detailed above for Q46C with the followingdifferences. Complementary mutagenic primers BB158 (5>AGGGATGGTTTCACACTTTTGGAACTG>3)(SEQ.D.NO. 71) and BB157 (5>CAG TTCCAAAAGTGTGAAACCATCCCT>3)(SEQ.ID.NO. 72), which change the GCT codon for A50 to a TGTcodon for cysteine, replaced BB154 and BB153 respectively in the primaryPCR reactions. The [BB125×BB158] and [BB126×BB157] PCR reactions gaveproducts of the expected sizes: ˜270 bp for [BB125×BB156] and ˜440 bpfor [BB126×BB155].

The substitution mutation D77C was constructed and sequence verifiedusing the protocols detailed above for Q5C with the followingdifferences. Complementary mutagenic primers BB138 (5>TAGGAGGGTCTCACACCAAGCAGCAGA>3)(SEQ.ID.NO. 73) and BB137(5>TCTGCTGCTTGGTGTGAGACCCTCCTA>3)(SEQ.ID.NO. 74), which change the GATcodon for D77 to a TGT codon for cysteine, replaced BB130 and BB129respectively in the primary PCR reactions. The [BB125×BB138] and[BB126×BB137] PCR reactions gave products of the expected sizes: ˜350 bpfor [BB125×BB138] and ˜345 bp for [BB126×BB137]. The product of thesecondary PCR reaction was “cleaned up” using the QIAquick PCRPurification (Qiagen), digested with Bgl II and Sal I (New EnglandBioLabs) according to the vendor protocols, “cleaned up” using theQIAquick PCR Purification Kit and run out on a 2% agarose gel. The ˜275bp Bgl II-Sal I fragment of interest was gel purified using a QIAEX IIGel Extraction Kit (Qiagen) according to the vendor protocol. Thisfragment was ligated with pBBT177 that had been cut with Bgl II and SalI, treated with calf intestinal alkaline phosphatase (New EnglandBioLabs) and gel purified. The ligation reaction was used to transformE. coli and plasmids from resulting transformants were sequenced toidentify a clone containing the D77C mutation and having the correctsequence throughout the ˜275 bp Bgl II-Sal I segment.

The substitution mutation T106C was constructed and sequence verifiedusing the protocols detailed above for D77C with the followingdifferences. Complementary mutagenic primers BB140(5>CAGGGGAGTCTCACACACCCCCACCCC>3)(SEQ.ID.NO. 75) and BB139(5>GGGGTGGGGGTGTGTGAGACTCCCCTG>3)(SEQ.ID.NO. 76), which change the ACAcodon for T106 to a TGT codon for cysteine, replaced BB138 and BB137respectively in the primary PCR reactions. The [BB125×BB140] and[BB126×BB139] PCR reactions gave products of the expected sizes: ˜435 bpfor [BB125×BB140] and ˜260 bp for [BB126×BB139].

The substitution mutation T108C was constructed and sequence verifiedusing the protocols detailed above for D77C with the followingdifferences. Complementary mutagenic primers BB142 (5>CTTCATCAGGGGACACTCTGTCACCCC>3)(SEQ.ID.NO. 78) and BB141(5>GGGGTGACAGAGTGTCCCCTGATGAAG>3)(SEQ. ID. NO. 79), which change the ACTcodon for T108 to a TGT codon for cysteine, replaced BB138 and BB137respectively in the primary PCR reactions. The [BB125×BB142] and[BB126×BB141] PCR reactions gave products of the expected sizes: ˜440 bpfor [BB125×BB142] and ˜250 bp for [BB126×BB141].

The substitution mutation Q101C was constructed and sequence verifiedusing the protocols detailed above for D77C with the followingdifferences. Complementary mutagenic primers BB162 (5>CACCCCCACCCCACATATCACACAGGC>3)(SEQ.ID.NO. 80) and BB161(5>GCCTGTGTGATATGTGGGGTGGGGGTG>3)(SEQ.ID.NO. 81), which change the CAGcodon for Q101 to a TGT codon for cysteine, replaced BB138 and BB137respectively in the primary PCR reactions. The primary reactions wereperformed in a Perkin-Elmer GeneAmp® PCR System 2400 thermal cycler. Thereaction program entailed: 95° C. for 5 minutes, 30 cycles of [95° C.for 30 seconds, 62° C. for 30 seconds, 72° C. for 1 minute] followed by72° C. for 7 minutes and a hold at 4° C. The [BB125×BB162] and[BB126×BB161] PCR reactions gave products of the expected sizes: ˜425 bpfor [BB125×BB162] and ˜275 bp for [BB126×BB161]. The secondary PCRreaction was also performed in a Perkin-Elmer GeneAmp® PCR System 2400thermal cycler. This reaction program entailed: 96° C. for 5 minutes, 25cycles of [95° C. for 30 seconds, 60° C. for 30 seconds, 72° C. for 1minute] and a hold at 4° C. Following digestion with Bgl II and Sal I,the products of this reaction were cleaned up using the QIAquick PCRPurification Kit but not gel purified prior to ligation.

The substitution mutation E107C was constructed and sequence verifiedusing the protocols detailed above for Q101C with the followingdifferences. Complementary mutagenic primers BB164 (5>CATCAGGGGAGTACATGTCACCCCCAC>3)(SEQ.ID.NO. 81) and BB163(5>GTGGGGGTGACATGTACTCCCCTG ATG>3)(SEQ.ID.NO. 82), which change the GAGcodon for E107 to a TGT codon for cysteine, replaced BB162 and BB161respectively in the primary PCR reactions. The [BB125×BB164] and[BB126×BB163] PCR reactions gave products of the expected sizes: ˜440 bpfor [BB125×BB164] and ˜255 bp for [BB126×BB163].

The substitution mutation C98S was constructed and sequence verifiedusing the protocols detailed above for Q101C with the followingdifferences. Complementary mutagenic primers BB160 (5>CCCCTGTATCACAGAGGMITCCAGGTC>3)(SEQ.ID.NO. 83) and BB159(5>GACCTGGAAGCCTCTGTGATACA GGGG>3)(SEQ.ID.NO. 84), which change the TGTcodon for C98S to a TCT codon for serine, replaced BB162 and BB161respectively in the primary PCR reactions. The [BB125×BB160] and[BB126×BB159] PCR reactions gave products of the expected sizes: ˜415 bpfor [BB125×BB160] and ˜285 bp for [BB126×BB159].

The cysteine substitution mutation S163C was constructed as follows. Themutagenic reverse oligonucleotide BB143(5>CGCGAATICTTATTCCTTACATCTTAAACT=TC>3)(SEQ.ID.NO. 85) was designed tochange the codon AGT for serine at position 163 to a TGT encodingcysteine and to span the nearby Eco RI site. This oligo was used in PCRwith the forward, non-mutagenic, primer BB125. A 50 μl PCR reaction wasperformed in 1× Promega PCR buffer containing 1.5 mM MgCl₂, each primerat 0.2 μM, each of dATP, dGTP, dTTP and dCTP at 200 μM, 1 ng of templateplasmid pBBT131 1.5 units of Taq Polymerase (Promega), and 0.25 units ofPfu Polymerase (Stratagene). Reactions were performed in a RobocyclerGradient 96 thermal cycler (Stratagene). The reaction program entailed:96° C. for 3 minutes, 25 cycles of [95° C. for 1 minute, 60° C. for 1.25minutes, 72° C. for 1 minute] followed by a hold at 6° C. A 5 μl aliquotof the PCR reaction was analyzed by agarose gel electrophoresis andfound to produce a single fragment of the expected size ˜610 bp. Theremainder of the reaction was “cleaned up” using the QIAquick PCRPurification (Qiagen) according to the vendor protocol, digested withSal I and Eco RI (New England BioLabs) according to the vendorprotocols, ethanol-precipitated, resuspended in 20 μl of 10 mM Tris-HClpH 8.5 and run out on a 2% agarose gel. The ˜42 bp Sal I-Eco RI fragmentof interest was gel purified using a QIAEX II Gel Extraction Kit(Qiagen) according to the vendor protocol. This fragment was ligatedwith pBBT132 that had been cut with Sal I and Eco RI, treated with calfintestinal alkaline phosphatase (New England BioLabs) and gel purified.The ligation reaction was used to transform E. coli and plasmids fromresulting transformants were sequenced to identify a clone containingthe E107C mutation and to have the correct sequence throughout the ˜42bp Sal I-Eco RI segment.

A mutation was also constructed that added a cysteine following thecarboxyterminal amino acid of the IFN-α2 coding sequence. This mutant,termed *167C was constructed using the protocols described above for theconstruction of the S163C mutant with the following differences. Themutagenic reverse oligonucleotide BB144(5>CGCGAATTCTtAACATTCCTTACTTCTTAAACTTTC>C>3)(SEQ.ID.NO. 86) which adds aTGT codon for cysteine between the GAA codon for E165 and the TAA stopcodon and spans the nearby Eco RI site was used in the PCR reaction inplace of BB143.

The substitution mutation E 165C was constructed and sequence verifiedusing the protocols detailed above for S163C with the followingdifferences. The mutagenic reverse oligonucleotide BB165 (5>CGCGAATTCTTAACACTTACTTCTTAAACT>3)(SEQ.ID.NO. 87) which changes the GAA codon forE165 to a TGT codon for cysteine and spans the nearby Eco RI site wasused in the PCR reaction in place of BB143. The PCR reaction wasperformed in a Perkin-Elmer GeneAmp® PCR System 2400 thermal cycler. Thereaction program entailed: 95° C. for 5 minutes, 30 cycles of [95° C.for 30 seconds, 62° C. for 30 seconds, 72° C. for 1 minute] followed by72° C. for 7 minutes and a hold at 4° C. Following digestion with Eco RIand Sal I, the products of this reaction were cleaned up using theQIAquick PCR Purification

For expression in E. coli as proteins secreted to the periplasmic space,the STII-IFN-α2 genes encoding the 17 muteins were excised from thepUC18-based pBBT177 derivatives as Nde 1-Eco RI fragments of 590 bp andsubcloned into the pCYB1 expression vector that had been used to expresswild type STII-IFNα-2. For expression experiments, these plasmids wereintroduced into E. coli coli W3110.

Using procedures similar to those described here, one can constructother cysteine muteins of IFN-α2. The cysteine muteins can besubstitution mutations that substitute cysteine for a natural aminoresidue in the IFN-α2 coding sequence, insertion mutations that insert acysteine residue between two naturally occurring amino acids in theIFN-α2 coding sequence, or addition mutations that add a cysteineresidue preceding the first amino acid, C1, of the IFN-α2 codingsequence or add a cysteine residue following the terminal amino acidresidue, E165, of the IFN-α2 coding sequence. The cysteine residues canbe substituted for any amino acid, or inserted between any two aminoacids, anywhere in the IFN-α2 coding sequence. Preferred sites forsubstituting or inserting cysteine residues in IFN-α2 are in the regionpreceding Helix A, the A-B loop, the B-C loop, the C-D loop, the D-Eloop and the region distal to Helix E. Other preferred sites are thefirst or last three amino acids of the A, B, C, D and E Helices.Preferred residues in these regions for creating cysteine substitutionsare D2, L3, P4, T6, H7, S8, Q20, R22, K23, S25, F27, S28, K31, D32, R33,D35, G37, F38, Q40, E41, E42, F43, G44, K49, T52, N65, S68, T69, K70,D71, S72, S73, A74, A75, D77, E78, T79, Y89, Q90, Q91, N93, D94, E96,A97, G102, V103, G104, V105, P109, M111, K112, E113, D114, S115, K131,E132, K133, K134, Y135, S136, A139, S152, S154, T155, N156, L157, Q158,E159, S160, L161, R162, and K164. Cysteine residues also can be insertedimmediately preceding or following these amino acids. Another preferredsite for adding a cysteine residue would be preceding C1, which we referto as *-1C.

One also can construct IFN-α2 muteins containing a free cysteine bysubstituting another amino acid for one of the naturally-occurringcysteine residues in IFN-α2. The naturally-occurring cysteine residuethat normally forms a disulfide bond with the substituted cysteineresidue is now free. The non-essential cysteine residue can be replacedwith any of the other 19 amino acids, but preferably with a serine oralanine residue. A free cysteine residue also can be introduced intoIFN-α2 by chemical modification of a naturally occurring amino acidusing procedures such as those described by Sytkowski et al. (1998).

Using procedures similar to those described in Examples 20-22, one canexpress the proteins in E. coli, purify the proteins, PEGylate theproteins and measure their bioactivities in an in vitro bioassay. TheIFN-α2 muteins also can be expressed in eukaryotic cells such as insector mammalian cells, using procedures similar to those described inExamples 16-20, or related procedures well known to those skilled in theart.

B. E. coli Expression of rIFN-α2 Cysteine Muteins

To assess expression, cultures of the rIFN-α2 muteins were grown andinduced as described above for wild type rIFN-α2. Typically, 45 mlcultures were grown in 250 ml shake flasks at 37° C. in a gyrotoryshaker water bath at ˜180-200 rpm. We observed that vigorous aeration ofshake flask cultures results in reduced levels of IFNα2 protein in thesupernatants of the osmotic shock lysates. Therefore we routinely usedconditions that were sub-optima for aeration but preferable for solublerIFN-α2 and rIFN-α2 mutein production. The induced cultures wereincubated for ˜16 h, harvested and subjected to osmotic shock asdetailed above for wild-type rIFN-α2 with the exception that cystine wasadded to a final concentration of 5 mM to the buffers used for theosmotic shock procedure. Adding cystine to the osmotic shock buffersresulted in significantly improved chromatographic properties for thefirst two muteins analyzed, Q5C and S163C. Interferon muteins nottreated with cystine consistently eluted from the S-Sepharose column asbroad bands, which, when analyzed by non-reducing SDS-PAGE, showedmultiple molecular weight species at and around the expected monomermolecular weight. These species most likely represent misfolded orincompletely folded interferon variants. In contrast, interferon muteinstreated with cystine during the osmotic shock procedure eluted from theS-Sepharose column as sharp bands, which, when analyzed by non-reducingSDS-PAGE, consisted of only one interferon species that co-migrated withthe interferon wild type standard. Recoveries of the purified interferonmuteins from the S-Sepharose column also were 1.5- to 2-fold greaterwhen cystine was included in the osmotic shock buffers. Based upon theseresults, 5 mM cystine was added to the osmotic shock buffers used topurify all the cysteine muteins.

SDS-PAGE analysis of the osmotic shock supernatants of the muteinsshowed most to have reduced (as compared to wild type) levels of the 19kDa rIFN-α2 band. Two muteins, Q5C and S163C, were expressed at levelsequivalent to wild type interferon, and several hundred micrograms ofeach of these muteins were readily purified as detailed below. Eightmuteins (C1S, 43C44, N45C, Q46C, Q48C, A50C, D77C and T108C) wereessentially undetectable in osmotic shock supernatants. The remainingseven muteins (F47C, C98S, Q101C, T106C, E107C, E165C, *166C) weredetected at varyingly reduced levels (as compared to wild type) inosmotic shock supernatants. Some of these muteins (T106C, C98S, E107C,and Q101C) were purified from osmotic shock supernatants, but only smallquantities of the pure proteins were recovered. Certain muteins, C98S,Q101C, T106C, E107C and *166C, were expressed at relatively high levelsbut accumulated primarily in an insoluble form, presumably in theperiplasm. These proteins comigrated with wild type rIFN-α2 standardsunder reducing conditions indicating that the STII leader had beenremoved. Qualitative assessments of relative expression levels of themuteins are summarized in Table 4.

C. Purification of rIFN-α2 Cysteine Muteins.

In order to purify the rIFN-α2 muteins, typically, a 325 ml culture in a2 liter shake flask, or a 500 ml culture in a 2 liter baffled shakeflask, were grown at 37° C. in a gyrotory shaker water bath (NewBrunswick Scientific) ˜170-220 rpm. Cultures were grown, induced,harvested, and subjected to osmotic shock as described in Example 20.Resulting osmotic shock supernatants were processed immediately orstored at −80° C.

The soluble IFN-α2 muteins in the osmotic shock supernatants werepurified using S-Sepharose and Cu⁺⁺ IMAC chromatography as detailedabove for purification of wild type rIFN-α2. All of the muteins testedbound tightly to the copper column and eluted under conditions similarto wild-type rIFN-α2. This result suggests that the conformations of thecysteine muteins are similar to that of native rIFN-α2, at least in theregions that comprise the metal-binding pocket.

Non-reducing SDS-PAGE analysis of the purified Q5C, C98S, Q101C, T106C,E107C, S163C, and *166C cysteine muteins showed that the muteins wererecovered predominantly as monomers, migrating at the expected molecularweight of ˜19 kDa. C98S migrated with a slightly higher molecular weightthan the other rINF-α2 muteins due to the absence of the nativeCys1-Cys-98 disulfide bond. Some of the purified muteins contained smallamounts of disulfide-linked rIFN-α2 dimers. The molecular weights of thedimer species were approximately 37-38 kDa.

D. Bioactivities of rIFN-α2 Cysteine Muteins.

Biological activities of the purified Q5C and S163C rIFN-α2 cysteinemuteins were measured in the Daudi growth inhibition assay described inExample 20. Protein concentrations were determined using Bradford or BCAprotein assay kits (Bio-Rad Laboratories and Pierce). Commercial wildtype rIFN-α2 and rIFN-α2 prepared by us were analyzed in parallel on thesame days to control for interday variability in the assays. The muteinsinhibited proliferation of Daudi cells to the same extent as the wildtype rIFN-α2 control proteins, within the error of the assay. The meanIC₅₀ for the Q5C mutein was 13 pg/ml which is similar to the mean IC₅₀sof the wild type rIFN-α proteins. The mean IC₅₀ for the S163C proteinwas 27 μg/ml. These data are summarized in Table 4. TABLE 4 Expressionand in vitro Bioactivities of IFN-α2 Cysteine Muteins RelativeExpression Total Percent Mean IC₅₀ IC₅₀ Range³ IFN-α2 Protein MutationLocation Cellular¹ Soluble² (pg/ml) (pg/ml) rIFN-α2⁴ — — — 16 +/− 7 8-29(n = 10) rIFN-α2⁵ — ++++ ˜33 13 +/− 4 7-19 (n = 10) C1S N-terminalregion⁶ +/− 0 Q5C N-terminal region ++++ ˜20 13 9, 11, 15, 18 43C44 A-Bloop ++ 0 N45C A-B loop ++ 0 Q46C A-B loop +/− 0 F47C A-B loop ++++ ˜5Q48C A-B loop +/− 0 A50C A-B loop +/− 0 D77C B-C loop +/− 0 C98SC-helix⁷ +++++ ˜5-10 Q101C C-D loop +++++ ˜5-10 T106C C-D loop +++++˜5-10 E107C C-D loop +++++ ˜5-10 T108C C-D loop +/− 0 S163C C-terminalregion ++++ ˜33 27 +/− 8 18-40 (n = 6) E165C C-terminal region +++ ˜20*166C C-terminus +++ ˜20¹Relative accumulation of the IFN-α2 protein in whole cell extracts²Portion of the IFN-α2 protein in the osmotic shock supernatant,estimated from SDS-PAGE gels³IC₅₀ values from individual experiments⁴Commercial wild type rIFN-α2 (Endogen, Inc.)⁵Wild type rIFN-α2 prepared by Bolder Biotechnology, Inc.⁶Mutation creates a free cysteine (C98) in the C-helix⁷Mutation creates a free cysteine (C1) in the N-terminal region

EXAMPLE 22 PEGylation, Purification and Bioactivity of PEG-Q5C andPEG-S163C

A. PEGylation of IFN-α Cysteine Muteins.

A small-scale PEGylation experiment was performed with the purifiedrIFN-α2 cysteine muteins to identify conditions that allowed theproteins to be monoPEGylated at the free cysteine residue.Over-reduction of the proteins was monitored by non-reducing SDS-PAGE,looking for a shift to a higher than expected apparent molecular weightas a result of protein unfolding, or for the appearance of multiplePEGylated species generated as the result of native disulfide reduction.Initial titration experiments were performed with the Q5C protein. Oneμg aliquots of purified Q5C were incubated with increasingconcentrations of TCEP [Tris(2-carboxyethyl) phosphine]-HCl at roomtemperature in 100 mM Tris, pH 8.5 in the presence of varying amounts ofexcess 5 kDa maleimide-PEG. After 60 min, the reactions were stopped andimmediately analyzed by non-reducing SDS-PAGE. The amounts of TCEP andPEG reagent that yielded significant amounts of monoPEGylated Q5Cprotein (molecular weight of approximately 28 kDa by non-reducingSDS-PAGE), without modifying wild type rIFN-α2, were used for furtherexperiments. The titration experiments indicated that a 10-fold molarexcess of TCEP and 20-fold excess of 5 kDa maleimide PEG gave around 60%monoPEGylated protein without detectable di or tri-PEGylated protein, ormodification of wild type rIFN-α2.

These conditions also were used to PEGylate several other rIFN-α2muteins. One μg aliquots of purified wild type and the rIFN-α2 muteins(Q5C, T106C, E107C, S163C) were incubated for 1 hour with a 10-foldmolar excess TCEP and a 20-fold molar excess of 5 kDA maleimide PEG atpH 8.5 at room temperature. The four muteins were monoPEGylated tovarying degrees (estimated to be from 30-60%) based on SDS-PAGE analysisof the reaction mixtures. Wild-type rIFN-α2 showed no detectablePEGylation under these conditions. Control experiments indicated thatthe Q5C, T106C, E107C and S163C cysteine muteins needed to be reducedwith TCEP to be PEGylated. These data indicate that the PEG molecule isattached to the cysteine residue introduced into the Q5C, T106C, E107Cand S163C proteins.

B. Preparation and Purification of PEG-Q5C IFN-α2 and PEG-163C:

Larger quantities of the Q5C and S163 muteins were PEGylated so thatbiological activities of the PEGylated proteins could be measured. Forthe Q5C protein, the PEGylation conditions used for the small-scaleexperiments were scaled to 140 μg protein to give sufficient materialfor purification and characterization. The larger PEGylation reactionwas performed for 1 hr at room temperature, diluted 10× with 20 mM MES,pH 5.0, adjusted to pH 3.0, and then loaded quickly onto an S-Sepharosecolumn using conditions similar to those described for initialpurification of the rIFN-α2 muteins. The presence of the PEG moietydecreased the protein's affinity for the resin, allowing the PEGylatedprotein to be separated from the non-PEGylated protein. The chromatogramfrom the S-Sepharose column showed two major protein peaks eluting atapproximately 190 mM NaCl and 230 mM NaCl. The early eluting major peak(eluting at approximately 190 mM NaCl) was determined to bemono-PEGylated Q5C by SDS-PAGE. The apparent molecular weight ofmonoPEgylated Q5C is approximately 28 kDa by SDS-PAGE. The later elutingmajor peak (eluting at approximately 230 mM NaCl) was determined to beunreacted Q5C protein. Fractions from the early eluting peak containingpredominantly PEG-Q5C, were pooled and used for bioactivitymeasurements.

The S163 C cysteine mutant was PEGylated at a 90 μg scale and purifiedusing protocols essentially identical to those described for PEG-Q5C.

C. Bioactivities of PEG-Q5C and PEG-S163C Cysteine Muteins:

Biological activity of the purified PEG-Q5C protein was measured in theDaudi cell assay described in Example 20. Concentration of the proteinwas determined using a Bradford dye binding assay. The PEG-Q5C proteinshowed a similar dose-response curve and reached the same level ofmaximal growth inhibition as wild type rIFN-α2 and the non-modified Q5Cprotein, within the error of the assay. The mean IC₅₀ for the PEG-Q5Cmutein was ˜22 pg/ml, which is within 2-fold of the IC₅₀ valuesdetermined for wild type IFN-α2 and the unmodified Q5C proteins analyzedon the same days (Table 5). Bioactivity of the PEG-Q5C protein issignificantly greater than that of rIFN-α2 that has been PEGylated withnon-specific, amine-reactive PEG reagents. The latter protein has anIC₅₀ of 164 pg/ml in the Daudi cell assay (Monkarsh et al., 1997). Thesedata are summarized in Table 5.

Bioactivity experiments also were performed with the PEG-S163C protein.The PEG-S163C protein also was biologically active and inhibited Daudicell proliferation to the same extent as wild type rIFN-α2, within theerror of the assay. The average IC₅₀ for the PEG-S163C protein was about42 pg/ml, which is better than the amine-Pegylated IFN-α. TABLE 5Bioactivity of PEG-Q5C IFN-α EC₅₀ Range¹ Mean EC₅₀ (pg/ml) IFN-α Protein(pg/ml) Exp A Exp B Exp C Endogen rIFN-α 13 9 11 20 rIFN-α² 12 10 10 16Q5C 12 8.5 11 18 PEG-Q5C 22 18 18 30 Amine-PEGylated-IFN-α³ 164 — — —¹Data from three experiments.²rIFN-α2 prepared by Bolder Biotechnology, Inc.³Data from Monkarsh et al. (1997)

EXAMPLE 23

In vivo efficacy of the PEGylated GH cysteine muteins can be tested inhypophysectomized HYPOX) rats. This is a well-characterized model of GHdeficiency (Cox et al., 1994; Clark et al., 1996). GH stimulates bodyweight gain and bone and cartilage growth in HYPOX rats (Cox et al.,1994; Clark et al., 1996). Hypophysectomized Sprague-Dawley rats can bepurchased from a commercial supplier such as Charles River (Wilmington,Mass.). Typically, rats are hypophysectomized between 40 and 50 days ofage and weigh approximately 120 g. Groups of 8 rats should receivesubcutaneous injections of rhGH, PEG-Cys-GH or placebo (vehiclesolution) at specified intervals and weight gain measured daily over a10 day period. Rats should be weighed daily at the same time per day toeliminate possible variables associated with feeding. In addition tooverall weight gain, bone growth (tibial epiphysis width) can bemeasured. At time of sacrifice, the right and left proximal tibialepiphyses can be removed and fixed in formalin. The fixed tibias can besplit at the proximal end in a saggital plane, stained with silvernitrate and exposed to a strong light (Greenspan et al., 1949). Thewidth of the cartilaginous epiphseal plate can be measured using astereomicroscope equipped with a micrometer eyepiece. Ten measurementsshould be made for each epiphysis and the means +/−SEM for the combinedvalues for the left and right tibias should be calculated. Comparisonsbetween groups can be made using a Students T test for singlecomparisons and one-way analysis of variance for multiple comparisons.P<0.05 should be considered significant.

Efficacy of the GH cysteine muteins modified with 10 kDa or 20 kDa PEGscan be tested by administering the proteins to the rats daily, everyother day, every third day, every fourth day or following a singleinjection Five μg of non-PEGylated hGH administered twice a day (10 μgBID) by subcutaneous injection gives a strong growth response in theHYPOX rat model (Cox et al., 1994; Clark et al., 1996). In initialexperiments different groups of rats should receive subcutaneousinjections of 0.08, 0.4, 2, 10, or 50 μg of the PEGylated Cys-GHproteins/injection/rat. Control rats should receive vehicle solutiononly. Additional control groups should receive non-PEGylated rhGH (10μg/BID) and 10 μg non-PEGylated hGH using the same dosing regimen as thePEGylated Cys-GH proteins. Administration of the PEGylated GH cysteinemuteins to the HYPOX rats should result in an increase in body weightgain and tibial epyphysis width growth compared to the vehicle-treatedgroup.

Efficacy of the PEGylated GH cysteine muteins also can be tested inrodent models of cachexia. Dexamethasone (DEX) can be administered tothe rats to induce weight loss. Groups of normal Sprague-Dawley rats(200-225 g) should receive daily subcutaneous injections ofdexamethasone (200 μg/rat, approximately 1 mg/kg). This amount ofdexamethasone should induce a loss of approximately 5-6 g over an 8 dperiod. Vehicle or varying doses of the PEGylated GH cysteine muteinscan be administered to the rats once, daily, every other day, everythird day or every fourth day in different experiments. Different groupsof rats should receive subcutaneous injections of 0.08, 0.4, 2, 10, or50 μg of the PEGylated Cys-GH proteins/injection/rat Additional controlsshould include a group of rats that will receive no DEX or injections, agroup of rats that receives DEX and non-PEGylated rhGH (10 μg BID) and agroup of rats that receives DEX and non-PEGylated rhGH (10 μg daily,every other day, every third day, or every fourth day, depending uponthe experiment, i.e., frequency that the PEGylated GH cysteine mutein isadministered). Animals should be weighed daily. Food and waterconsumption should be monitored daily. At time of sacrifice, internalorgans should be weighed. Statistical analyses should be performed asdescribed for the HYPOX rat studies. Animals treated with the PEGylatedGH cysteine muteins should lose less weight than the vehicle-treatedanimals.

In vivo efficacy of the PEGylated EPO cysteine muteins can be measuredin normal rats by demonstrating that the proteins stimulate increases inhemocrit and erythropoiesis compared to vehicle-treated animals. EPOstimulates a significant increase in hematocrit in rats when dosed on adaily basis (Matsumoto et al., 1990; Vaziri et al., 1994; Baldwin etal., 1998). Sprague-Dawley rats can be purchased from a commercialsupplier such as Charles River (Wilmington, Mass.). Groups of 5 ratsshould receive subcutaneous injections of BV rEPO, PEGylated EPOcysteine mutein or placebo (vehicle solution) at specified intervals forup to five days. On day 6 the animals should be sacrificed and bloodsamples collected for hematocrit and complete blood cell count (CBC)analysis, which can be performed by a commercial laboratory.Hematopoietic tissues (liver and spleen) should be collected, weighedand fixed in formalin for histopathologic analyses to look for evidenceof increased erythropoiesis. Bone marrow should be removed from variouslong bones and the sternum for unit particle preparations andhistopathologic analysis to look for evidence of increasederythropoiesis. Comparisons between groups should be made using aStudents T test for single comparisons and one-way analysis of variancefor multiple comparisons. P<0.05 should be considered significant. ThePEGylated EPO cysteine muteins should stimulate increases in hematocritand erythropoiesis in the rats compared to the vehicle-injected animals.Efficacy of the PEGylated EPO cysteine muteins modified with 10 kDa or20 kDa PEGs can be tested when administered once, every other day orevery third day. 100 IU/kg (˜800 ng/kg) of non-PEGylated EPOadministered once per day (160 ng SID/200 g rat) by subcutaneousinjection gives a significant increase in hematocrit (Matsumoto et al.,1990; Vaziri et al., 1994; Baldwin et al., 1998). In initial experimentsdifferent groups of rats should receive subcutaneous injections of 0.32,1.6, 8, 40 or 160 ng of the PEGylated EPO cysteine muteins. Control ratsshould receive vehicle solution only. Additional control groups shouldreceive non-PEGylated rEPO (160 ng/SID) and 160 ng non-PEGylated rEPOusing the same dosing regimen as the PEGylated EPO cysteine muteins.

Efficacy of the PEGylated EPO muteins also can be tested inchemotherapy-induced anemia models. Cisplatin-induced anemia is awell-characterized rodent model of chemotherapy-induced anemia and hasdirect relevance to the human clinical setting. rEPO reverses the anemiain this model when administered at daily doses of 100 Units/kg(Matsumoto et al., 1990; Vaziri et al., 1994; Baldwin et al. 1998).Sprague-Dawley rats (˜200 g) should be treated on day 0 with anintraperitoneal injection of Cisplatin (3.5 mg/kg) to induce anemia andrandomized to various treatment groups. Efficacy of the PEGylated EPOcysteine muteins modified with 10 kDa or 20 kDa PEGs can be tested whenadministered once (on day 1), every other day or every third day.Different groups of rats should receive subcutaneous injections of 0.32,1.6, 8, 40 or 160 ng/injection of the PEGylated EPO cysteine muteins.Rats should be injected with the test compounds for up to 8 days. Onecontrol group of rats should receive daily subcutaneous injections ofrEPO (100 Units/kg). Another control group should not receive theinitial Cisplatin injection but should receive injections of salineusing the same dosing schedules used for the PEGylated EPO cysteinemuteins. On day 9 the rats should be sacrificed and blood and tissuesamples obtained for comprehensive CBC and histopathology analyses. ThePEGylated EPO cysteine muteins should stimulate increases in hemocritand erythropoiesis in the rats compared to the vehicle-injected controlgroup.

IFN-α biological activity is relatively species-specific, which limitsthe range of preclinical animal models that can be studied. One modelthat can be used to measure in vivo efficacy of PEGylated IFN-α2cysteine muteins is inhibition of human tumor xenograft growth inathymic nude mice. Human IFN-α2 is not active on mouse cells andinhibition of human tumor xenograft growth in nude mice occurs through adirect antiproliferative effect on the human tumor cells. IFN-α2inhibits growth of a variety of primary human tumor xenografts and humantumor cell lines in athymic mice (Balkwill et al., 1985; Balkwill, 1986;Johns et al., 1992; Lindner and Borden, 1997). The primary endpoint, forthe studies should be tumor volume in treated mice. We expect to findthat the administration of the PEGylated IFN-α2 cysteine muteinsinhibits tumor growth (as measured by tumor volume) in the mice relativeto vehicle-treated animals. Athymic nude mice can be purchased from acommercial vendor such as Charles River. Each mouse should be injectedwith 2×10⁶ NIH-OVCAR-3 or MCF-7 tumor cells (the cell lines areavailable from the American Type Culture collection) on day 0 andrandomly assigned to test groups, consisting of ten mice each The tumorcells should be injected into the dermis overlying the mammary glandnearest the axillae. The different test groups should receivesubcutaneous injections of varying doses of wild type rIFN-α2, 10kDa-PEGylated IFN-α2 cysteine mutein, 20 kDa-PEGylated IF-α2 cysteinemutein or placebo (vehicle solution) at specified intervals: every day(SID), every other day (EOD) or every third day (ETD). Tumor volumesshould be determined at 4 day intervals by measuring the length andwidth of the tumors with calipers, as described by Linder and Borden(1997). At time of sacrifice, the tumors should be excised and weighed.Mean tumor volumes +/−SEM for each test group should be calculated foreach sampling point. Comparisons between groups should be made using aStudents T test for single comparisons and one-way analysis of variancefor multiple comparisons. Five μg of non-PEGylated IFN-α2 administeredonce per day by subcutaneous injection inhibits growth of NIH-OVCAR-3cells and MCF-7 cells in athymic mice by 80% after 6 weeks (Lindner andBorden, 1997). Either cell line can be used for these studies. TheNIH-OVCAR-3 line (available from the ATCC) does not require estrogen forgrowth, as do the MCF-7 cells. Xenograft experiments with MCF-7 cellsrequire that the mice be oophorectomized and implanted with estrogenpellets (Lindner and Borden, 1997). In initial experiments, differentgroups of mice should receive subcutaneous injections of 1 or 5 μg perinjection of rIFN-α2, 10-kDa-PEGylated IFN-α2 cysteine mutein or20-kDa-PEGylated IFN-α2 cysteine mutein using every day, every other dayor every third day dosing schedules. Dosing should begin on day 2following injection of the tumor cells into the mice. Control miceshould receive vehicle solution only. In the every other day and everythird day dosing experiments, an additional positive control groupshould receive daily subcutaneous injections of 5 μg unmodified rIFN-α2.

REFERENCES

-   Abdel-Meguid, S. S., Shieh, H -S., Smith W. W., Dayringer, H. E.,    Violand, B. N. and Bentle, L. A. (1987) Proc. Natl. Acad. Sci. USA    84: 6434-6437.-   Abrahmsen, L., Moks, T., Nilsson, B. and Uhlen, M. (1986) Nucleic    Acids Res. 14:7487-7500.-   Abuchowski, A., Kazo, G. M., Verhoest, C. R., van Es, T., Kafkewitz,    D., Nucci, M. L., Viau, A. T. and Davis, F. F. (1984) Cancer    Biochem. Biophys. 7: 175-186.-   Alt, F. W., Kellems, R.E., Bertino, J. R., and    Schimke, R. T. (1978) J. Biol. Chem. 253:1357-1370.-   Ayres, M. D., Howard, S. C., Kuzio, J., lopez-Ferber, M., and    Possee, R. D. (1994) Virology 202:586-605.-   Barik, S. (1993) in “Methods in Molecular Biology”, White, B. A.,    ed. (Humana Press, Totawa, N.J.), 15: 277-286.-   Bazan, F. (1991) Immunology Today 11: 350-354.-   Bazan, J. F. (1992) Science 257: 410-411.-   Becker, G. W. and Hsiung, H. M. (1986) FEBS Lett. 204: 145-150.-   Bewley et al., (1969) Biochem. 8: 4701-4708.-   Bill, R. M., Winter, P. C., McHale, C. M., Hodges, V. M., Elder, G.    E., Caley, J., Flitsch, S. L., Bicknell, R. and    Lappin, T. R. J. (1995) Biochem. Biophys. Acta 126: 35-43.-   Blatt, L. M., Davis, J. M. Klein, S. B. and Taylor, M.W. (1996) J.    Interferon and Cytokine Research 16: 489-499.-   Boissel, J. -P., Lee, W. -R., Presnell, S. R., Cohen, F. E. and    Bunn, H. F. (1993) J. Biol. Chem. 268:15983-15993.-   Bollag, D. M., Rozycki, M. D. and Edelstein, S. J. (1996) Protein    Methods, 415 pages, Wiley-Liss, NY, N.Y.-   Braxton, S. M. (1998) U.S. Pat. No. 5,766,897.-   Chamow & Ashkenazi (1996), Trends in biotech 14:52-60.-   Chang, C. N., Rey, B., Bochner, B., Heyneker, H. and Gray, G. (1987)    Gene: 189-196.-   Cheah, K -C., Harrison, S., King, R., Crocker, L., Wells, J. R. E.    and Robins, P. (1994) Gene, 138: 9-15.-   Clark, R., Olson, K., Fuh, G., Marian, M., Mortensen, D., Teshimna,    G., Chang, S., Chu, H., Mukku, V., Canova-Davis, E., Somers, T.,    Cronin, M., Winkler, M. and Wells, J. A. (1996) J. Biol. Chem. 271:    21969-21977.-   Cox, G. N. and McDermott, M. J. (1994) WO 9412219.-   Cox, G. N., McDermott, M. J., Merkel, E., Stroh, C. A., Ko, S. C.,    Squires, C. H., Gleason, T. M. and Russell, D. (1994) Endocrinology    135: 1913-1920.-   Cox, G. N. and Russell, D. (1994) WO 9,422,466.-   Crouse, G. F., McEwan, R. N., and Pearson, M. L. (1983) Mol. Cell.    Biol. 3:257-266.-   Cunningham, B. C. and Wells, J. (1989) Science 244: 1081-1085.-   Cunningham, B. C., Jhurani, P., Ng, P. and Wells, J. A. (1989)    Science 243: 1330-1336.-   Cunningham, B. C., Ultsch, M., de Vos, A. M., Mulkerrin, M. G.,    Clauser, K. R. and Wells, J. A. (1991) Science 254: 821-825.-   Daopin, S., Piez, K. A., Ogawa, Y., and Davies, D. R. (1992) Science    257: 369-373. Davis, S., Aldrich, T. H., Stahl, N., Pan, L., Taga,    T., Kishimoto, T., Ip, N. Y. and Yancopoulus, G. D. (1993) Science    260: 1805-1808.-   Davies, A. H. (1995) Curr. Opin. Biotechnol. 6:543-547.-   DeChiara, T. M., Erlitz, F. and Tarnowski, k., S. J. (1986) Methods    Enzymology, 119: 403-15.-   de la Llosa, P., Chene, N. and Maral, J. (1985) FEBS Letts. 191:    211-215.-   Denefle, P., Kovarik, S., Ciora, T. Gosselet, M., Benichou, J. -C.,    Latta, M. Guinet, F., Ryter, A. and Mayaux, J. -F. (1989) Gene 85,    499-510.-   de Vos, A. M., Ultsch, M. and Kossiakof, A. A. (1992) Science 255:    306-312.-   Diederichs, K., Boone, T. and Karplus, A. (1991) Science 154:    1779-1782.-   Evinger, M. and Pestka, S. (1981) Methods Enzymol. 79:362-368.-   Fuh, G., Cunningham, B. C., Fukunaga, R., Nagata, S., Goeddel, D. V.    and Wells, J. A. (1992) Science 256: 1677-1680.-   Fujimoto, K., Fukuda., T., and Marumoto., R. (1988) J. Biotechnol.    8:77-86.-   Geisse, S., Gram, H. Kleuser, B. and Kocher, H. P. (1996) Prot.    Express. Purif. 8:271-282.-   Goeddel, D. V., Heyneker, H. L., Hozumi, T., Arentzen, R., Itakura,    K., Yansura, D. G., Ross, M. J., Miozzari, G., Crea, R. and    Seeburg, P. H. (1979) Nature 281: 544-548.-   Goodson, R. J. and Katre, N. V. (1990) Biotechnology 8: 343-346.-   Greenspan, F. S., Li., C. H., Simpson, M. E. and Evans, H. M. (1949)    Endocrinology 45:455-463.-   Ghrayeb, I., Kimura, R., Takahara, M., Hsiung, H., Masui, Y. and    Inouye, M. (1984) EMBO J. 3:2437-2442.-   Hannum, C., Culpepper, J., Campbell D., McClanahan, T., Zurawski, S.    et al. (1994) Nature 368: 643-648-   Hershfield, M. S., Buckley, R. H., Greenberg, M. L. et    al., (1987) N. Engl. J. Medicine 316: 589-596.-   Hill, C. P., Osslund, T. D. and Eisenberg, D. (1993) Proc. Natl.    Acad. Sci. USA 90:5167-5171.-   Horisberger, M. A. and Di Marco, S. (1995) Pharmac. Ther. 66:    507-534.-   Horoszewicz, J. S., Leong, S. S. and Carter, W. A. (1979) Science    206:1091-1093.-   Horton, R. M. (1993) in “Methods in Molecular Biology”, White, B.    A., ed. (Humana Press, Totawa, N.J.), v. 15, 214-250.-   Hsiung, H. M., Mayne, N. G. and Becker, G. W. (1986) Biotechnology    4: 991-995.-   Imai, N., Kawamura, A., Higuchi, M., Oh-eda, M., Orita, T.,    Kawaguchi, T., and Ochi, N. (1990) J. Biochem. 107:352-359.-   Innis, M. A., Gelfand, D. H., Sninsky, J. J. and White, T. J.    eds. (1998) “PCR Protocols: A Guide to Methods and Applications”    (Academic Press, San Diego, Calif.).-   Jacobs, K., Shoemaker, C., Rudersdorf, R., Neill, S. D., Kaufman, R.    J., Mufson, A., Seehra, J., Jones, S. S., Hewick, R., Fritsch, E.    F., Kawakita, M., Shimizu, T. and Miyake, T. (1985) Nature 313:    806-810.-   Johnson, D. L., Middleton, S. A., McMahon, F. Barbone, F. P., Kroon,    D., Tsao, E., Lee, W. H., Mulcahy, L. S. and Jolliffe, L. K. (1996)    Protein Expression Purif. 7:104-113.-   Kadonaga, J., Gautier, A. Straus, D. R., Charles, A. D., Edge, M. D.    and Knowles, J. R. (1984) J. Biol. Chem. 259: 2149-2154.-   Khan, F. R. and Rai, V. R. (1990) Bioprocess Technology 7:161-169.-   Karasiewicz, R., Nalin, C. and Rosen, P. (1995) U.S. Pat. No.    5,382,657.-   Katre, N. V. (1990) J. Immunology 144: 209-213.-   Katre, N. V., Knauf M. J. and Laird, W. J. (1987) Proc. Natl. Acad.    Sci. USA 84: 1487, 1491.-   Kaufman, R. J. (1990) Meth. Enzymol. 185:537-566.-   Kingsley, D. M. (1994) Genes Dev. 8: 133-146.-   Kinstler, O. B., Gabriel, N. E., Farrar, C. E. and    DePrince, R. B. (1996) International Patent Application Number WO    96/11953.-   Knauf, M. J., Bell, D. P., Hirtzer, P., Luo, Z. -P., Young, P. D.    and Katre, N. V. (1988) J. Biol. Chem. 263: 15064-15070.-   Komatsu, N., Nakauchi, H., Miwa, A., Ishilara, T., Eguchi, M.,    Moroi, M., Okada, M., Sato, Y., Wada, H., Yawata, Y., et al., (1991)    Cancer Research 51: 341-348.-   Koshland, D. and Botstein, D. (1980) Cell 20: 749-760.-   Kutty G., Kutty, R. K., Samuel, W., Duncan, T., Jaworski, C., and    Wiggert, B. (1998) Biochem. Biophys. Res. Commun. 246: 644-649.-   Lawton, L. N., Bonaldo, M. F., Jelenc, P. C., Qiu, L., Baumes, S.    A., Marcelino, R. A., de Jesus, G. M., Wellington, S., Knowles, J.    A., Warburton, D., Brown, S., and Soares, M. B. (1997) Gene 203:    17-26.-   Li, C. H. (1982) Mol. Cell. Biochem. 46: 31-41.-   Lin, F. -K., Suggs, S., Lin, C. -H., Browne, J. K., Smalling, R.,    Egrie, J. C., Chen, K. K., Fox, G. M., Martin, F., Stabinsky, Z.,    Badrawi, S. M., Lai, P. -H. and Goldwasser, E. Proc. Natl. Acad.    Sci. USA 82: 7580-7584.-   Lu, H. S., Boone, T. C., Souza, L. M., and Lai, P. H. (1989) Arch.    Biochem. Biophys. 268: 81-92.-   Lucas, B. K., Giere, L. M., DeMarco, M. A., Shen, A., Chisolm, V.,    and Crowley, C. W. (1996) Nucleic Acids Res. 24:1774-1779.-   Lydon, N. B., Favre, C., Bove, S., Neyret, O., Benureau, S.,    Levine, A. M., Seelig, G. F., Nagabhushan, T. L. and    Trotta, P. P. (1985) Biochemistry 24: 4131-4141.-   MacGillivray, M. H., Baptista, J. and Johnson, A. (1996) J. Clin.    Endocrinol. Metab. 81: 1806-1809.-   Maisano, F., Testori, S. A., and Grandi, G. (1989) J. Chromatograph.    472: 422-427.-   Mark, D. F., Lin, L. S. and Lu, S -D. Y. (1985) U.S. Pat. No.    4,518,584.-   Mark, D. F., Lu, S. d., Creasey, A. a., Yamamoto, R. and    Lin, L. S. (1984) Proc. Natl. Acad. Sci. USA 81: 5662-5666.-   Martal, J., Chene, N. and de la Llosa, P. (1985) FEBS. Letts. 180:    295-299.-   Martin, F. H, Suggs, S. V., Langley, K. E., Lu, X. S., Ting, J.,    Okino, K. H., Morris, F. C., McNiece, I. K., Jacobsen, F. W.,    Mendiaz, E. A., Birkett, N. C. et al., (1990) Cell 63: 203-211.-   Massague, J. (1990) Annu. Rev. Cell Biol. 6: 597-641.-   Matthews, D. J., Topping, R. S., Cass, R. T. and    Giebel, L. B. (1996) Proc. Natl. Acad. Sci. USA 93: 9471-9476.-   McDonald, N. Q. and Hendrickson, W. A. (1993) Cell 73: 421-424.-   McKay, D. B. (1992) Science 257:412.-   Meyers, F. J., Paradise, C., Scudder, S. A., Goodman, G. and    Konrad, M. (1991) Clin. Pharmacol Ther. 49: 307-313.-   Milburn, M. V., Hassell, A. M., Lambert, M. H., Jordan, S. R.,    Proudfoot, A. E., Graber, P. and Wells, T. N. C. (1993) Nature 363:    172-176.-   Mills, J. B., Kostyo, J. L., Reagan, C. R., Wagner, S. A.,    Moseley, M. H. and Wilhelm, A. E. (1980) Endocrinology 107: 391-399.-   Mockridge, J. W., Aston, R., Morrell, D. J. and Holder, A. T. (1998)    Eur. J. Endocrin. 138: 449-459.-   Monkarsh, S. P., Ma, Y., Aglione, A., Bailon, P. et al. (1997) Anal.    Biochem 247: 434-440.-   Mordenti, J., Chen, S. A., Moore, J. A., Ferrailo, B. L. and    Green, J. D. (1991) Pharmacol. Res. 8:1351-1359.-   Morehead, H., Johnson, P. D. and Wetzel, R. (1984) Biochemistry 23:    2500-2507.-   Morioka-Fujimoto, K., Marumoto, R. and Fukuda, T. (1991) J. Biol.    Chem. 266: 1728-1732.-   Mott, H. R. and Campbell, I. D. (1995) Current Opinion in Structural    Biology 5: 114-121.-   Ostermeier, M., De Sutter., K., Georgiou, G. (1996) J. Biol. Chem.    271: 10616-10622.-   Paonessa, G., Graziani, R., de Serio, A., Savino, R., Ciapponi, L.,    Lahm, A., Ssalvati, A. L., Toniatti, C. and Ciliberto, G. (1995)    EMBO J. 14: 1942-1951.-   Paul, W. E. ed. (1989) “Fundamental Immunology” (Raven Press, New    York, N.Y.).-   Pestka, S., Langer, J. A., Zoon, K. C. and Samuel, C. E. (1987) Ann.    Rev. Biochem 56: 727-777.-   Picken, R. N., Mzaitis, A. J., Maas, W. K., Rey, M. and    Heyneker, H. (1983) Infect and Immun. 42: 269-275.-   Powers, R., Garrett, D. S., March, C. L, Frieden, E. A,    Gronenbor, A. M. and Clore, G. M. (1992) Science 256: 1673-1677.-   Ranjan, A. and Hasnain, S. E. (1995) Virus Genes 2:149-153.-   Redfield, C., Smith L. J., Boyd, J., Lawrence, G. M. P., Edwards, R.    G., Smith, R. A. G., and Dobson, C. M. (1991) Biochemistry 30:    11029-11035.-   Roitt, I. M., Brostoff, J., and Male, D. K. eds. (1989) “Immunology”    (Gower Medical Publishers, New York, N.Y. and London, UK)-   Rowlinson, S. W. Barnard, R., Bastias, S., Robins, A. J.,    Brinkworth, R. and Waters, M. J. (1995) J. Biol. Chem. 270:    16833-16839.-   Shaw, G., Veldman., G. and Wooters, J. L. (1992) U.S. Pat. No.    5,166,322.-   Sytkowski, A. J., Lunn, E. D., Davis, K. L., Feldman, L. and    Siekman, S. (1998) Proc. Natl. Acad. Sci. USA 95: 1184-1188.-   Tanaka, H., Satake-Ishikawa, R., Ishikawa, M., Matsuki, S. and    Asano, K. (1991) Cancer Research 51: 3710-3714.-   Teh, L. -C. and Chapman, G. E. (1988) Biochem. Biophys. Res. Comm.    150: 391-398.-   Thompson, S. A. (1992) J. Biol. Chem. 267: 2269-2273.-   Trill, J. J., Shatzman, A. R., and Ganguly, S. (1995) Curr. Opin.    Biotechnol. 6:553-560.-   Tuma, R., Rosendahl, M. and Thomas, G. (1995) Biochem. 34:    15150-15156.-   Urlaub, G. and Chasin, L. A. (1980) Proc. Natl. Acad. Sci. USA    77:4216-4220.-   Van Den Berg, C. L., Stroh, C., Hilsenbeck, S. G., Weng, C. -N.,    McDermott, M. J., Cox, G. N. and Yee, D. (1997) Eur. J. Cancer 33:    1108-1113.-   Voss, T., Falkner, E., Ahorn, H. Krystek, E. Maurer-Fogy, I. Bodo,    G., Hauptmann, R. (1994) Biochem. J. 298, 719-725.-   Walter, M. R., Cook, W. J., Ealick, S. E., Nagabhusan, T. L.,    Trotta, P. T. and Bugg, C. E. (1992) J. Mol. Biol. 224: 1075-1085.-   Wang, G. L. and Semenza, G. L. (1993) Blood 82:3610-3615.-   White, B. A. (1993) in Methods in Molecular Biology, Vol. 15: PCR    Protocols: Current Methods and Applications edited by Humana Press,    Inc., Totowa, N. J.-   Wojchowski, D. M., Orkin, S. H. and Sytkowshi, A. J. (1987) Biochim.    Biophys. Acta 910: 224-232.-   Wrighton, N. C., Farrell, F. X., Chang, R. et al., (1996) Science    273: 458-463.-   Yamaoka, T., Tabata, Y. and Ikada, Y. (1994) J. Pharm. Sci. 83:    601-606.-   Baldwin, M. D., Zhou, X. J., Ing, T. S. and Vaziri, N. D. (1998)    ASAIO J. 44: 44-47.-   Balkwill, F. R., Goldstein, L. and Stebbing, N. (1985) Int. Cancer    35: 613-617.-   Balkwill, F. R. (1986) Methods Enzymology 119: 649-657.-   Cecil, R. and McPhee, J. R. (1959). Advances in Protein Chemistry    14, 255-389.-   Cox, G. N., McDermott, M. J., Merkel, E., Stroh, C. A., Ko, S. C.,    Squires, C. H., Gleason, T. M. and Russell, D. (1994) Endocrinology    135: 1913-1920.-   Greenspan, F. S., Li., C. H., Simpson, M. E. and Evans, H. M. (1949)    Endocrinology 45: 455-463.-   Johns, T. G., Mackay, I. R., Callister, K. A., Hertzog, P. J.,    Devenish, R. J. and Linnane, A. W. (1992) J. Natl. Cancer Institute    84: 1185-1190.-   Lindner, D. J. and Borden, E. C. (1997). Interferon and Cytokine    Research 17:681-693.-   Matsumoto, T., Endoh, K., Kamisango, K., Akamatsu, K., Koizumi, K.,    Higuchi, M., Imai, N., Misui, H and Kawaguchi, T. (1990) Br. J.    Haematol. 75: 463-468.-   Torchinskii, Y. M. (1971) in “Sulfhydryl and Disulfide Groups of    Proteins” in Studies of Soviet Science (1971) Nauka Press, Moscow.-   Trotta, P. B. (1986) Seminars in Oncology XIII Supplement 2: 3-12.-   Vaziri, N. D., Zhou, X. J., and Liao, S. Y. (1994) Am. J. Physiol.    266: F360-366.

While various embodiments of the present invention have been describedin detail, it is apparent that modifications and adaptations of thoseembodiments will occur to those skilled in the art. It is to beexpressly understood, however, that such modifications and adaptationsare within the scope of the present invention.

1-31. (Cancelled)
 32. An isolated, monoPEGylated erythropoietin proteinor derivative thereof, wherein said monoPEGylated erythropoietin proteincontains N24, N38 and N83 and wherein said monoPEGylated erythropoietinprotein comprises a polyethylene glycol attached to an amino acidlocated in a region of erythropoietin selected from the group consistingof: the A-B loop, the B-C loop, the C-D loop, the last three amino acidsin helix A, the first three or last three amino acids in helix B, thefirst three or last three amino acids in helix C, the first three orlast three amino acids in helix D, and the region preceding helix A,wherein said monoPEGylated erythropoietin protein has an EC₅₀ of lessthan about 1000 ng/ml, as measured in vitro by proliferation of a cellline that proliferates in response to erythropoietin.
 33. The isolated,monoPEGylated erythropoietin protein or derivative thereof of claim 32,wherein said monoPEGylated erythropoietin protein has an EC₅₀ of lessthan about 100 ng/ml.
 34. The isolated, monoPEGylated erythropoietinprotein or derivative thereof of claim 32, wherein said monoPEGylatederythropoietin protein has an EC₅₀ of less than about 10 ng/ml.
 35. Theisolated, monoPEGylated erythropoietin protein or derivative thereof ofclaim 32, wherein said monoPEGylated erythropoietin protein has an EC₅₀of less than about 1 ng/ml.
 36. The isolated, monoPEGylatederythropoietin protein or derivative thereof of claim 32, wherein saidmonoPEGylated erythropoietin protein contains less than 5% unPEGylatederythropoietin protein.
 37. The isolated, monoPEGylated erythropoietinprotein or derivative thereof of claim 32, wherein said PEGylatederythropoietin protein has an apparent molecular weight of greater thanabout 70 kDA, as measured by size exclusion column chromatography. 38.The isolated, monoPEGylated erythropoietin protein or derivative thereofclaim 32, wherein said polyethylene glycol has a molecular mass ofbetween 2 kDA and 40 kDA.
 39. The isolated, monoPEGylated erythropoietinprotein or derivative thereof of claim 32, wherein said polyethyleneglycol is selected from the group consisting of a linear polyethyleneglycol and a branched polyethylene glycol.
 40. The isolated,monoPEGylated erythropoietin protein or derivative thereof of claim 32,wherein said polyethylene glycol is a cysteine-reactive polyethyleneglycol.
 41. The isolated, monoPEGylated erythropoietin protein orderivative thereof of claim 40, wherein said polyethylene glycol isselected from the group consisting of: a maleimide-polyethylene glycol,a vinylsulfone-polyethylene glycol and an iodoacetyl-polyethyleneglycol.
 42. The isolated, monoPEGylated erythropoietin protein orderivative thereof of claim 32, wherein said monoPEGylatederythropoietin protein comprises at least one added sulfur groupattached to at least one amino acid residue in said erythropoietinprotein.
 43. The isolated, monoPEGylated erythropoietin protein orderivative of claim 42, wherein said polyethylene glycol is attached tosaid added sulfur group in said erythropoietin protein.
 44. Theisolated, monoPEGylated erythropoietin protein or derivative thereof ofclaim 42, wherein said added sulfur group is attached to a lysineresidue in said erythropoietin protein.
 45. The isolated, monoPEGylatederythropoietin protein or derivative of claim 44, wherein saidpolyethylene glycol is attached to said added sulfur group in saiderythropoietin protein.
 46. The isolated, monoPEGylated erythropoietinprotein or derivative thereof of claim 32, wherein said monoPEGylatederythropoietin protein is glycosylated.
 47. The isolated, monoPEGylatederythropoietin protein or derivative thereof of claim 46, wherein atleast one amino acid selected from the group consisting of N24, N38 andN83 is glycosylated and wherein at least one amino acid selected fromthe group consisting of N24, N38 and N83 is not glycosylated.
 48. Theisolated, monoPEGylated erythropoietin protein or derivative thereof ofclaim 47, wherein at least one amino acid selected from the groupconsisting of N38 and N83 is glycosylated and wherein N24 is notglycosylated.
 49. The isolated, monoPEGylated erythropoietin protein orderivative thereof of claim 47, wherein at least one amino acid selectedfrom the group consisting of N24 and N83 is glycosylated and wherein N38is not glycosylated.
 50. The isolated, monoPEGylated erythropoietinprotein or derivative thereof of claim 47, wherein at least one aminoacid selected from the group consisting of N24 and N38 is glycosylatedand wherein N83 is not glycosylated.
 51. A mixture comprising two ormore of the monoPEGylated erythropoietin proteins or derivative thereofof claim
 32. 52. The isolated, monoPEGylated erythropoietin protein orderivative thereof of claim 32, wherein a polyethylene glycol isattached to an amino acid at a position selected from the groupconsisting of: 1, 2, 3, 4, 5, 8, 25, 26, 27, 28, 30, 31, 32, 34, 36, 39,40, 43, 44, 45, 47, 49, 50, 52, 53, 54, 55, 57, 58, 77, 78, 79, 84, 85,86, 88, 89, 107, 110, 111, 113, 114, 115, 116, 117, 118, 120, 121, 122,123, 124, 125, 126, 127, 128, 131, and 132 of erythropoietin.
 53. Theisolated, monoPEGylated erythropoietin protein or derivative thereof ofclaim 32, wherein said monoPEGylated erythropoietin protein comprises apolyethylene glycol attached to an amino acid located in the A-B loop oferythropoietin.
 54. The isolated, monoPEGylated erythropoietin proteinor derivative thereof of claim 53, wherein a polyethylene glycol isattached to the amino acid at position 26 of erythropoietin.
 55. Theisolated, monoPEGylated erythropoietin protein or derivative thereof ofclaim 32, wherein said monoPEGylated erythropoietin protein comprises apolyethylene glycol attached to an amino acid located in the B-C loop oferythropoietin.
 56. The isolated, monoPEGylated erythropoietin proteinor derivative thereof of claim 32, wherein said monoPEGylatederythropoietin protein comprises a polyethylene glycol attached to anamino acid located in the C-D loop of erythropoietin.
 57. The isolated,monoPEGylated erythropoietin protein or derivative thereof of claim 56,wherein a polyethylene glycol is attached to the amino acid at position126 of erythropoietin.
 58. The isolated, monoPEGylated erythropoietinprotein or derivative thereof of claim 32, wherein said monoPEGylatederythropoietin protein comprises a polyethylene glycol attached to oneof the last three amino acids in helix A.
 59. The isolated,monoPEGylated erythropoietin protein or derivative thereof of claim 32,wherein said, monoPEGylated erythropoietin protein comprises apolyethylene glycol attached to one of the first three or one of thelast three amino acids in helix B.
 60. The isolated, monoPEGylatederythropoietin protein or derivative thereof of claim 32, wherein saidmonoPEGylated erythropoietin protein comprises a polyethylene glycolattached to one of the first three or one of the last three amino acidsin helix C.
 61. The isolated, monoPEGylated erythropoietin protein orderivative thereof of claim 32, wherein said monoPEGylatederythropoietin protein comprises a polyethylene glycol attached to oneof the first three or one of the last three amino acids in helix D. 62.The isolated, monoPEGylated erythropoietin protein or derivative thereofof claim 32, wherein said monoPEGylated erythropoietin protein comprisesa polyethylene glycol attached to an amino acid located in the regionpreceding helix A of erythropoietin.
 63. The isolated, monoPEGylatederythropoietin protein or derivative thereof of claim 32, wherein apolyethylene glycol is attached to amino acid 1 of erythropoietin.
 64. Amethod for treating a condition treatable with erythropoietin,comprising administering to a patient a composition comprising theerythropoietin protein or variant thereof of claim 32.